pathophysiology of persistent doxorubicin cardiotoxicity · universidade de coimbra faculdade de...

Universidade de CoimbraFaculdade de Ciências e TecnologiaDepartamento de Ciências da Vida

Pathophysiology of persistentdoxorubicin cardiotoxicity

Metabolic landscaping of the epigenome

Author:André Ferreira

Supervisors:Paulo J. Oliveira

António J. Moreno

June, 2015

André Ferreira: Pathophysiology of persistent doxorubicin cardiotoxicity, Metaboliclandscaping of the epigenome, B.Sc. Biology, June 2015

This work was performed at the Center for Neuroscience and Cell Biology, De-partment of Life Sciences, University of Coimbra, Portugal, under the supervisionof Dr. Paulo J. Oliveira (CNC, UC) and Dr. António J. Moreno (DCV, UC). Dr.Teresa Cunha-Oliveira also supervised the present work.

The work presented in this dissertation was funded by FEDER funds throughthe Operational Programme Competitiveness Factors – COMPETE and nationalfunds by the Portuguese Foundation for Science and Technology (FCT) –, byan institutional grant UID/NEU/04539/2013 to the CNC and by QREN project“Stemcell based platforms for Regenerative and Therapeutic Medicine” [#4832,reference CENTRO-07-ST24-FEDER-002008] funded through FEDER/QREN 2007–2013. Also supported by a PhD fellowship from FCT to Filipa S. Carvalho(SFRH/BD/64694/2009) and by research grants to Dr. Paulo J. Oliveira (PTDC/DTP-FTO/1180/2012) and Dr. António Moreno (PTDC/SAU-OSM/104731/2008).

Acknowledgements

Firstly I would like to thank my supervisor Dr. Paulo Oliveira for introducing meto this project and without whose help this work could not have been possible. Iwould also like to thank Dr. Teresa Oliveira for her continuous advice, guidance,and most of all for her endless patience and time spent training me in most ofwhat I currently know.

I would like to acknowledge the help and time given by all the people inthe MitoXT group – past and present. The making of this thesis would havebeen a diabolical endeavor had it not been for the moments, minutes or hours ofassistance given whenever the need for help arose. Many thanks in particular forthe multiple times when someone volunteered to finish my experiments or takecare of my mess every time I was running late for my ride back home.

v

Abstract

Doxorubicin (DOX), a potent and broad-spectrum antineoplastic agent, causesa cumulative dose-dependent cardiomyopathy of late-onset that may ultimatelylead to congestive heart failure. The mechanisms responsible for the delayednature of DOX cardiotoxicity remain poorly understood. Since DOX induces apersistent depression in cardiomyocyte gene expression patterns, it is reasonableto hypothesize that a “toxicity memory” is being formed by means of epigeneticmodulation. To test this hypothesis, eight week-old male Wistar rats (n = 6 pergroup) were administered 7 weekly injections with DOX (2 mg kg−1) or equivalentvolume of saline solution and sacrificed two weeks after the last injection. Weassessed gene expression patterns by qPCR, global DNA methylation by ELISAand proteome lysine acetylation status by Western blot in cardiac tissue fromsaline and DOX-treated rats. We show for the first time that DOX treatmentdecreases global DNA methylation in a cardiac-specific fashion. These differenceswere accompanied by alterations in mRNA expression of multiple functional genegroups. Doxorubicin disrupted cardiac mitochondrial biogenesis, as demonstratedby reduced mtDNA levels and altered transcript levels for multiple mitochondrialgenes encoded by both nuclear and mitochondrial genomes. Transcription ofgenes involved in lipid metabolism and epigenetic modulation were also disturbed.Western blotting analyses indicated a differential protein acetylation pattern incardiac mitochondrial fractions of DOX-treated rats. Additionally, DOX treatmentincreased the activity of histone deacetylases. Overall, the results presented inthis thesis support previous observations showing that DOX impairs cardiacmitochondrial function and induces a switch in substrate metabolism. Theseresults are also consistent with the interplay between mitochondrial dysfunctionand epigenetic alterations in the persistent nature of DOX-induced cardiotoxicity.One possibility is that DOX may modulate gene expression patterns via alterationsto the epigenetic landscape, which could be related to a disturbance in metaboliteavailability.

Keywords: doxorubicin; epigenetics; heart; mitochondria; toxicology.

vii

Resumo

A doxorubicina (DOX), um potente fármaco antineoplástico com um amploespectro de acção, causa uma cardiomiopatia cumulativa, dose-dependente que emúltima instância pode levar a insuficiência cardíaca. Os mecanismos responsáveispor esta condição ainda não são conhecidos. Como a DOX induz uma depressãopersistente nos padrões de expressão genética em cardiomiócitos, podemos teorizarque a DOX imprime uma “memória tóxica” por meio de modulação epigenética.Para testar esta hipótese, ratos machos Wistar Han com oito semanas de idade(n = 6 por grupo) receberam oito injecções subcutâneas semanais de DOX(2 mg kg−1) ou de um volume equivalente de solução salina, sendo sacrificadosduas semanas após a última injecção. O nosso trabalho identificou padrõesde expressão genética por qPCR, metilação global de DNA por ELISA e opadrão de acetilação de lisinas no proteoma por Western blot em tecido cardíaco.Mostramos pela primeira vez que a DOX diminui os níveis de metilação globalde DNA no tecido cardíaco. Este efeito foi acompanhado por alterações naexpressão de mRNAs de vários conjuntos funcionais de genes. A DOX alterou abiogénese mitocondrial cardíaca, como identificado por uma redução dos níveis demtDNA e alterações dos níveis de transcriptos de múltiplos genes mitocondriaiscodificados por ambos os genomas (nuclear e mitocondrial). A transcrição degenes envolvidos no metabolismo de lípidos e em modulação epigenética tambémfoi alterada. Análises por Western blot revelaram um padrão de acetilaçãoproteica diferente em fracções mitocondriais de animais tratados com DOX.Adicionalmente, o tratamento com DOX aumentou a actividade enzimática dedeacetilases de histonas. Colectivamente, os resultados apresentados nesta tesesuportam observações anteriores que mostram que a DOX perturba a funçãomitocondrial e induz uma substituição no substrato metabólico preferencial nocoração, sendo também consistentes com a nossa hipótese de que uma acçãorecíproca entre disfunção mitocondrial e alterações epigenéticas é um factorimportante na natureza persistente da toxicidade induzida pela DOX. Uma possívelexplicação é a de que a DOX modula os padrões de expressão genética atravésde alterações na paisagem epigenética, o que pode ser consequência de umaperturbação na disponibilidade de metabolitos.

Palavras chave: coração; doxorubicina; epigenética; mitocôndria; toxicologia.

ix

Table of Contents

Abstract vii

Resumo ix

List of Figures xiii

List of Tables xv

List of Acronyms and Abbreviations xvii

1 Introduction 11.1 Doxorubicin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

1.1.1 Antitumor activity . . . . . . . . . . . . . . . . . . . . . . 21.1.2 Cardiotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . 4

1.2 Epigenetics, and the bioenergetics thereof . . . . . . . . . . . . . . 71.2.1 Histone acetylation . . . . . . . . . . . . . . . . . . . . . . 81.2.2 DNA methylation . . . . . . . . . . . . . . . . . . . . . . . 11

1.3 Objectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14

2 Materials and Methods 172.1 Animals and treatment protocol . . . . . . . . . . . . . . . . . . . 172.2 Subcellular fractionation . . . . . . . . . . . . . . . . . . . . . . . 172.3 DNA extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . 202.4 RNA extraction and cDNA synthesis . . . . . . . . . . . . . . . . 202.5 HAT activity assay . . . . . . . . . . . . . . . . . . . . . . . . . . 212.6 HDAC activity assay . . . . . . . . . . . . . . . . . . . . . . . . . 212.7 Enzyme-linked immunosorbent assay (ELISA) of global DNA m5C

content . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212.8 Primer design . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222.9 Quantitative real-time PCR . . . . . . . . . . . . . . . . . . . . . 222.10 Mitochondrial DNA integrity . . . . . . . . . . . . . . . . . . . . 262.11 Western blotting . . . . . . . . . . . . . . . . . . . . . . . . . . . 272.12 Densitometric analysis . . . . . . . . . . . . . . . . . . . . . . . . 272.13 Statistical analysis . . . . . . . . . . . . . . . . . . . . . . . . . . 28

xi

3 Results 293.1 Body, heart and liver weight . . . . . . . . . . . . . . . . . . . . . 293.2 DOX alters mitochondrial biogenesis transcriptomics and mitoepige-

nomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293.3 DOX alters transcription of acetyl CoA metabolism and transport

proteins and changes mitochondrial protein acetylation patterns . 323.4 DOX alters the transcription and activity of histone modulators . 323.5 DOX alters SAM metabolism transcripts and decreases global DNA

m5C levels in cardiac tissue . . . . . . . . . . . . . . . . . . . . . 34

4 Discussion 374.1 Mitochondrial biogenesis . . . . . . . . . . . . . . . . . . . . . . . 374.2 Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394.3 Epigenetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41

5 Conclusions 43

A Supplementary Information 45

References 58

xii

List of Figures

1.1 Chemical structure of Doxorubicin . . . . . . . . . . . . . . . . . 21.2 Formation of ROS through DOX redox cycling . . . . . . . . . . . 41.3 Conrad Waddington’s visual metaphor for the epigenetic landscape 71.4 Acetylation and deacetylation cycle of histone lysine residues . . . 81.5 Overview of the citrate and the carnitine shuttles . . . . . . . . . 91.6 DNA methylation mechanism . . . . . . . . . . . . . . . . . . . . 121.7 Simplified overview of folate-mediated one-carbon metabolism . . 131.8 Relationship between cellular metabolism and epigenetic modifications 15

2.1 Timeline of the treatment protocol. . . . . . . . . . . . . . . . . . 182.2 Schematic workflow of the subcellular fractionation protocol . . . 19

3.1 Effect of DOX treatment on body mass gain . . . . . . . . . . . . 303.2 DOX compromises mitochondrial biogenesis transcriptomics and

mitoepigenomics . . . . . . . . . . . . . . . . . . . . . . . . . . . 313.3 DOX alters the transcripts of genes involved in the metabolism and

availability of Ac-CoA in cardiac tissue, but does not change totallysine acetylation patterns . . . . . . . . . . . . . . . . . . . . . . 33

3.4 DOX alters transcripts and activity of histone modifiers . . . . . . 343.5 DOX decreases transcripts involved in the metabolism of SAM and

decreases global m5C content in cardiac tissue . . . . . . . . . . . 35

A.1 Representative immunoblots of Western blot experiments showingrelative levels of PGC-1α in nuclear fractions and mitochondrialfractions extracted from cardiac tissue . . . . . . . . . . . . . . . 45

xiii

List of Tables

2.1 List of oligonucleotide primer sequences used for qPCR . . . . . . 23

3.1 Impact of DOX treatment on body, heart and liver weights in rats 29

xv

List of Acronyms andAbbreviations

5-MTHF 5-Methyl-tetrahydrofolate

Ac-CoA Acetyl coenzyme AACL ATP citrate lyase

CAT Carnitine acetyltransferaseCACT Carnitine–acylcarnitine translocasecDNA Complementary DNACPT1B Carnitine palmitoyltransferase I BCyt c Cytochrome c

DNMT DNA methyltransferaseDOX DoxorubicinDTT Dithiothreitol

ELISA Enzyme-linked immunosorbent assayETC Electron transport chain

FFA Free fatty acid

GSH Glutathione, reduced

HAT Histone acetyl transferaseHcy HomocysteineHDAC Histone deacetylase

m5C 5-MethylcytosineMAT Methionine adenosyltransferasemtDNA Mitochondrial DNA

nDNA Nuclear DNANRF Nuclear respiratory factor

OxPhos Oxidative phosphorylation

xvii

PGC Peroxisome proliferator-activated receptor gamma coactivatorPPAR Peroxisome proliferator-activated receptor

qPCR Quantitative real-time polymerase chain reaction

ROS Reactive oxygen species

SAH S -Adenosyl-l-homocysteineSAL SalineSAM S -Adenosyl-l-methionineSTM Sucrose-Tris-magnesium

TBS-T Tris-buffered saline with TweenTFAM Mitochondrial transcription factor ATHF Tetrahydrofolate

xviii

Chapter 1

Introduction

1.1 DoxorubicinDoxorubicin (Adriamycin®, DOX) was one of the first anthracyclines∗ to beisolated from strains of Streptomyces actinobacteria in the 1960s (Arcamone et al.,1969). Since its clinical introduction in the 1970s, DOX has remained one of themost important components in several currently used chemotherapy protocolsfor treating breast, ovarian and gastric carcinomas, sarcomas, leukemias, non-Hodgkin’s and Hodgkin’s lymphoma, multiple myeloma and many other cancers(Sterba et al., 2013; Simunek et al., 2009). The impact of anthracycline-basedtherapies in the field of oncology is particularly noteworthy in pediatric oncology,where the 5-year survival rate for childhood cancer has increased from around30 % in the 1960s to over 70 % in the modern era, which explains why over 50 % ofchildhood cancer survivors are estimated to have received anthracycline treatment(Sterba et al., 2013; Simunek et al., 2009).

Figure 1.1 shows the chemical structure of DOX. It contains the basic structureof all anthracyclines, which is a tetracyclic anthraquinone-containing aglyconelinked to an aminosugar, and is distinguished mainly by a short side chain with acarbonyl group at C-13, a hydroxyl group at C-14 and the aminosugar daunosamineattached by a glycosidic bond to the C-7 of the tetracyclic ring. Because theanthracycline template has so much potential for modification, more than 2000structural variants of this class of compounds have been produced and tested forantitumor activity in an effort to find more effective and safer chemotherapeuticagents (Krohn, 2008). None of them have shown any significant advantage overpreviously existing compounds, and thus only a small number of anthracyclinesare currently employed in clinical settings – DOX and Daunorubicin, the firstanthracyclines to be isolated, being among the most relevant ones (Krohn, 2008).

Despite having over 40 years of extensive clinical utilization, DOX’s mechanismof action remains a matter of controversy to this day, as novel mechanisms arecontinuously being proposed. The cytostatic and cytotoxic actions of DOX incancer cells have been attributed to various mechanisms, most often: 1) DNA

∗Anthracyclines are defined as red–orange dyes containing a skeleton of 7,8,9,10-tetrahydro-tetracene-5,12-quinone with mono- to tetrasaccharide moieties attached, usually to ring D ofthe aglycone (Laatsch and Fotso, 2008).

1

2 Introduction

3

4

A

O

5

B

O

6

C

OH

7

D

O

O

OHNH2

8

9

OH

13

O

14

OH1011

OH

12

O

1

2

Figure 1.1. Chemical structure ofDoxorubicin.

intercalation, 2) topoisomerase II inhibition, 3) generation of free radicals withconsequent induction of oxidative stress and 4) apoptosis induction – althoughprobably as an outcome of the aforementioned events (Gewirtz, 1999). Novelalternative mechanisms recently proposed include inhibition of DNA methylationenzymes (Yokochi and Robertson, 2004) and nucleosome destabilization inducedby chromatin torsional stress (Yang et al., 2014). This diversity of antitumormechanisms is likely to underlie the broad spectrum of activity displayed by DOX.

As is the case for every anticancer agent, DOX is a double-edged sword dueto its toxic side effects in healthy tissue – cardiac muscle tissue in particular.Administration of DOX commonly results in relatively harmless short-term acuteeffects such as nausea, diarrhea, alopecia and electrocardiography alterations thatare frequently reversible (Sterba et al., 2013; Carvalho et al., 2009). Chroniccardiotoxic effects, on the other hand, are of much greater concern. Cumulativedoses exceeding 500–550 mg/m2 result in a clinically unacceptable risk of devel-oping congestive heart failure (CHF) (Lefrak et al., 1973), thus severely limitingthe clinical utility of this compound. The onset of this cardiomyopathy may bedelayed until as many as 20 years after cessation of treatment (Steinherz et al.,1991).

The mechanisms by which this cardiotoxicity arises appear to be very differentfrom the ones responsible for DOX’s antitumor activity. A plausible consequenceof this observation is the possibility of developing cardioprotective treatmentprotocols that can be performed in parallel with DOX-based chemotherapeuticprotocols withouth diminishing the drug’s efficacy.

1.1.1 Antitumor activityAs previously mentioned, DOX’s antitumor activity is mostly atributted to variousDNA-related interactions, as would be expected given the reliance of rapidlyproliferating tumor cells on DNA integrity.

DNA intercalation

Some of the earliest studies addressing the antitumor activity of anthracyclinesreported an inhibition of both DNA and RNA synthesis (Di Marco et al., 1965;

1.1 Doxorubicin 3

Bremerskov and Linnemann, 1969) and a high affinity of anthracyclines for DNA(Calendi et al., 1965). Naturally, DNA binding was promptly suggested as themechanism underlying the antitumor activity of anthracyclines. Indeed, DNA-anthracycline interactions have been confirmed by numerous sources of evidenceover time, from X-ray diffraction studies (Quigley et al., 1980; Nunn et al., 1991)to in silico analyses (Box, 2007).

The chemical structure of anthracyclines makes them suitable DNA interca-lating agents. Under the double helical structure, DNA can accommodate theplanar tetracyclic ring between neighboring base pairs. This DNA-anthracyclinecomplex is further stabilized by electrostatic interactions between the negativelycharged DNA phosphate backbone and the positively charged amino group ofthe aminosugar (Box, 2007; Beretta and Zunino, 2008). DNA intercalation isknown to occur at clinically relevant concentrations of DOX (Coldwell et al., 2008),however only a very small fraction of total DOX participates in this process (Yanget al., 2014). DNA intercalation is therefore unlikely to be the most significantmediator of DOX’s antitumor activity.

Topoisomerase II poisoning

DNA topoisomerases are enzymes responsible for removal of knots and tanglesgenerated during nuclear processes such as DNA replication and transcription.Topoisomerase II in particular is of critical importance, since its activity is abso-lutely required for chromosome segregation and therefore cell cycle progression(Nitiss, 1998). One step in the catalytic cycle of this enzyme consists in theintroduction of a short-lived double-strand DNA break that allows for the re-laxation of supercoils, thereby restoring functional DNA topology (Champoux,2001). A covalent enzyme-DNA complex – referred to as the “cleavable complex”– is established during this step, so as to safely sequester, process and repairthe topoisomerase II-mediated double-stranded DNA breaks (Wilstermann andOsheroof, 2003). Many anthracyclines (DOX included) belong to the class of drugsknown as topoisomerase II poisons, which stabilize the cleavable complex andprevent the DNA religation step (Larsen et al., 2003). Increasing the lifetime ofthis complex enhances the likelihood of incorrectly resolving the DNA breaks via,for example, illegitimate recombination events. Moreover, if left unresolved, DNAbreaks will trigger the DNA damage response and ultimately lead to apoptosis.

Oxidative stress

Generation of free radicals by DOX can occur through multiple pathways. Dox-orubicin is a suitable substrate for many flavoenzymes that catalyze the oxidationof NAD(P)H, such as the cytosolic eNOS (Vasquez-Vivar et al., 1997), the en-doplasmic reticulum and nuclear envelope cytochrome P450 reductase (Bachuret al., 1977; Bachur et al., 1982) and the mitochondrial NADH dehydrogenase(complex I) (Davies and Doroshow, 1986). The one-electron reduction of DOXinitiates a futile redox cycle (depicted in Fig. 1.2) that generates superoxide anions(O•−

2 ). Superoxide can then inactivate Fe−S cluster-containing enzymes (Flintet al., 1993), releasing in the process ferrous iron (Fe2+) that will in turn assist in

4 Introduction

O O OH O

O

OHNH2

OH

O

OH

OHO

FpH2NAD(P)H + H+ O•–2

O O• OH O

O

OHNH2

OH

O

OH

OHO−

FpNAD(P)+ O2

Fe2+

Fe3+

H2O2

OH– + •OH

SOD

DoxorubicinQuinone form

DoxorubicinSemiquinone form

Figure 1.2. Formation of ROS through DOX redox cycling. The quinone form of DOXaccepts an electron from either NADH or NADPH, in a reaction catalized by flavoprotein-containing enzymes such as NADH dehydrogenase (complex I) or cytochrome P450reductase. The resulting semiquinone radical cycles back to its quinone form upontransfering an electron to O2, thereby generating O•−

2 . Additional ROS can be formedin subsequent reactions, including the highly reactive •OH.

the iron-catalyzed Haber–Weiss reaction, leading to the formation of the highlyreactive hydroxyl radical (•OH) (Halliwell and Cross, 1994). Another suggestedmechanism is the formation of DOX-Fe complexes, in which the cycling betweenFe2+ and Fe3+ begets superoxide and hydroxyl radicals (Simunek et al., 2009).Cancer cells appear to demonstrate increased steady-state levels of oxidative stresswhen compared to normal cells (Aykin-Burns et al., 2009), and therefore are likelyto be more susceptible to apoptosis induction via increased reactive oxygen species(ROS) generation.

1.1.2 CardiotoxicityA variety of mechanisms have been proposed to be responsible for the developmentof DOX-induced cardiotoxicity. Mitochondrial interactions are some of the mostimplicated and best described mechanisms, and will be explored in greater detail

1.1 Doxorubicin 5

below.

Mitochondria-dependent toxicity

Cardiac muscle possesses one of the highest mitochondrial density values (ca.30 % cellular volume) of all tissues in the organism (Barth et al., 1992), reflectingits reliance on mitochondrial metabolism for energy production. Indeed, car-diomyocytes obtain over 90 % of their ATP through oxidative phosphorylation(OxPhos) (Ventura-Clapier et al., 2004). As such, the heart is a vulnerable organto drugs that target mitochondria, as is the case of DOX. Mitochondrial ultra-structural alterations were in fact one of the earliest features to be identified inDOX cardiotoxicity (Friedman et al., 1978; Aversano and Boor, 1983).

Doxorubicin has been shown to preferentially accumulate in cardiac tissue(Peters et al., 1981). The driving force behind this uptake is likely related to theelevated mitochondrial content displayed by this tissue. Doxorubicin binds withhigh affinity to cardiolipin (Goormaghtigh et al., 1980), a unique phospholipidfound almost exclusively in the inner mitochondrial membrane, and is retained inmitochondria as a result (Nicolay et al., 1986). Consequently, DOX is brought toclose proximity of the electron transport chain (ETC), facilitating the occurrenceof the aforementioned redox cycling in complex I. Moreover, cardiolipin-proteininteractions are essential for proper assembly and function of ETC proteins.Formation of DOX-cardiolipin complexes disrupts these interactions, possiblyleading to increased rates of electron leak (and thus, ROS formation) from theETC (Schlame et al., 2000).

If mitochondria are a major site of DOX-induced ROS formation, it followsthat mitochondria should also be a prime target for oxidative stress. This shouldbe especially the case for cardiac mitochondria, as cardiac tissue displays relativelylow levels of antioxidant defenses when compared to other organs including theliver (Doroshow et al., 1980). Consistent with this idea are the findings that DOXtreatment results in several oxidative stress markers in cardiac mitochondria, suchas lipid peroxidation (Mimnaugh et al., 1985), protein carbonylation (Ascensaoet al., 2006; Suliman et al., 2007) and mitochondrial DNA (mtDNA) oxidation(Palmeira et al., 1997; Serrano et al., 1999). Studies using transgenic animalmodels overexpressing mitochondrial exclusive (MnSOD) and non-exclusive (Gpx1)antioxidant enzymes have shown beneficial effects in DOX-induced cardiotoxicity(Yen et al., 1999; Xiong et al., 2006), further reinforcing the role of oxidative stressas a major culprit.

Not surprisingly, many of the expected functional consequences of mitochon-drial dysfunction have been reported for this pathology. Lipid peroxidation –particularly of the acyl chains in cardiolipin – not only compromises ETC function,as already mentioned, but also contributes to apoptosis. Cytochrome c (Cyt c),one of the ETC proteins that exists in close association with cardiolipin, has apivotal role in the intrinsic pathway of apoptosis, where its translocation from theintermembrane space to the cytosol is one of the required events (Raha and Robin-son, 2001). It has previously been shown that Cyt c translocation is a two-stepprocess, requiring both the permeabilization of the outer mitochondrial membraneand the detachment of Cyt c from the inner membrane (Ott et al., 2002). This

6 Introduction

last step occurs as a result of cardiolipin peroxidation, which compromises itsability to interact with Cyt c. Loss of mitochondrial calcium homeostasis and theconcurrent activation of the mitochondrial permeability transition are also likelyto contribute to apoptotic signaling (Solem et al., 1994; Zhou et al., 2001a). Boththe release of Cyt c and the ensuing activation of apoptosis have been confirmedin cardiomyocytes of DOX-treated rats (Childs et al., 2002). Finally, impairmentof overall OxPhos metabolism has also been extensively documented in murinemodels, as shown by the inhibition of ETC protein activity (Santos et al., 2002;Chandran et al., 2009), decreased state 3 respiration (oxygen consumption associ-ated with ATP synthesis) (Solem et al., 1994; Santos et al., 2002), increased state4 respiration (basal oxygen consumption) (Ji and Mitchell, 1994) and decreasedATP/ADP ratio (Oliveira et al., 2004).

Persistent cardiotoxicity

All the above notwithstanding, a satisfying explanation for the continued de-velopment of cardiotoxicity is yet to be found. It is clear from the progressivedevelopment of cardiomyopathy that DOX somehow imprints cardiomyocyteswith what can be considered a “toxic memory”. This notion is supported by oneof the few studies that employed a delayed model of DOX toxicity in vivo. Theauthors reported a sustained increase in oxidative stress after a recovery timeof 5 weeks – greater than the increase observed after 1 week of recovery (Zhouet al., 2001b) – suggesting that DOX imprinted a ROS-generating mechanismthat is not directly mediated by the drug itself. One possible mechanism that hasbeen suggested to be involved in this toxic memory is the oxidation of mtDNA(Berthiaume and Wallace, 2007a). Indeed, DOX was reported to induce a varietyof persistent insults to mtDNA in both animal models (Lebrecht et al., 2003)and human patients (Lebrecht et al., 2005). However, based on a number ofobservations, we consider that the possibility of long-term epigenetic modulationis a hypothesis worthy of investigation. Results from global gene expression arraysreveal that gene expression patterns – the endpoint of epigenetic modulation – arealtered in rats for up to 5 weeks after DOX treatment (Berthiaume and Wallace,2007b). The study reported a curious trend in metabolic genes: genes involvedin fatty acid oxidation were found to be downregulated while genes related toglycolysis were upregulated. The physiological implication of these findings waslater substantiated by 13C-isotopomer analyses (Carvalho et al., 2010), whichconfirmed the switch from the preferential oxidation of fatty acids to a moreglycolytic metabolism. Collectively, these results suggest that DOX induces along-term metabolic remodeling in cardiac tissue by acting at the transcriptionallevel, as opposed to the aforementioned short- to medium-term effects on OxPhosmetabolism that are dependent on the presence of the drug. Both the observedalterations in the transcriptome and the previously alluded to “toxic memory” con-cept would be consistent with DOX-induced persistent alterations in the epigeneticlandscape.

1.2 Epigenetics, and the bioenergetics thereof 7

Figure 1.3. Conrad Waddington’s vi-sual metaphor for the epigenetic land-scape. The immature cell depicted atthe top of the landscape is offered a setof paths towards potential differentiationoutcomes. Initial inputs determine whichpath will be followed, but a change incourse is possible upon overcoming a cer-tain threshold. Figure from Waddington(1957).

1.2 Epigenetics, and the bioenergetics thereofThe concept of epigenetics dates back to when DNA was yet to be identified asthe molecule of inheritance. In 1942, Conrad Waddington defined epigeneticsas “the branch of biology which studies the causal interactions between genesand their products, which bring the phenotype into being”. This concept wasWaddington’s attempt at bridging the gap between genetic information andobservable phenotypes that could not be accounted for by Mendelian inheritance.To this day, Waddington’s depiction of what he called the “epigenetic landscape”(Fig. 1.3) remains an adequate analogy for the concept.

Modern definitions of epigenetics refer to post-translational and chemicalmodifications of chromatin that do not alter the genomic sequence itself (Bergeret al., 2009). ∗ Chromatin is composed of repeating units of nucleosomes, eachconsisting of a 147 bp DNA segment wound around a histone octamer composedof two copies of each of the four core histones proteins H2A, H2B, H3 and H4(Luger and Hansen, 2005). Structurally, the core histones are mostly globular,with the exception of unstructured extensions protruding from the nucleosomecore that are known as “histone tails”. Chromatin post-translational modificationsoccur at the level of histones – particularly in the tail regions of histones H3and H4 (Kouzarides, 2007; Bannister and Kouzarides, 2011). Post-translationalmodifications described thus far include acetylation, methylation, phosphoryla-tion, ubiquitylation, sumoylation (SUMO: small ubiquitin-like modifier), ADPribosylation, deimination and proline isomerization (Kouzarides, 2007). Themajority of histone post-translational modifications are dynamic, and in normalconditions are regulated to adequately fit particular cellular contexts. Chemicalmodifications of chromatin, in turn, occur at the level of DNA, mainly in CpGdinucleotides. These include DNA methylation and hydroxymethylation (Guibertand Weber, 2013). Collectively, epigenetic modifications regulate the regionalarchitecture of chromatin, thereby controlling the accessibility of DNA to proteinssuch as transcription factors and ultimately determining gene expression patterns.Epigenetic modifications, in turn, are regulated by at least three factors: theexpression levels of chromatin-modifying enzymes, signaling pathways that recruit

∗Many authors further consider noncoding RNA-mediated chromatin regulation (Collinset al., 2011; Rinn and Chang, 2012) and transvection (Pirrotta, 1999) – a phenomenon thatrefers to interchromosomal interactions between homologous alleles – as epigenetic mechanisms.

8 Introduction

. . .

NH

⊕NH3

C

O

. . .

Lysine

. . .

NH

NHC

CH3

O

C

O

. . .

Acetyl-lysine

HATs

H3CC

O

SCoA HSCoA

HDACs

H3CC

O

OH

H2O

Figure 1.4. Acetylation and deacetylation cycle of histone lysine residues. Acetylmoieties are transfered from Ac-CoA to the ε-amino group of a lysine residue byHATs. The reverse reaction is performed by HDACs, which catalyze the hydrolysis ofacetyl-lysine to yield lysine and acetate. Sirtuins can also catalyze the deacetylation ofacetyl-lysine, although using a different catalytic mechanism from the one depicted here.

or displace these enzymes and the levels of substrates and cofactors required fortheir function (Lu and Thompson, 2012).

This section will limit the discussion to histone acetylation and DNA methyla-tion, along with the metabolic requirements of both processes. Potential routes ofinterference by DOX will also be addressed.

1.2.1 Histone acetylationHistone proteins have a positively charged surface owing to the abundant pres-ence of basic amino acids (lysine and arginine), specially so in the tail regions.Conversely, DNA is a mostly negative molecule due to its negatively charged phos-phate backbone. The resulting strong electrostatic interactions between histonesand DNA lead to the tight packing of DNA inside the nucleus. Although DNAcompaction is an absolute logistical requirement for eukaryotic cells (the unwoundhuman genome is about 2 m long), it also imposes accessibility constraints on thetranscriptional machinery. To circumvent this limitation, histone acetylation hasevolved so as to allow the spatial and temporal regulation of chromatin unpacking,thus enabling the transcriptional machinery to access specific DNA templates.

Acetylation of histones is a reaction catalyzed by histone acetyl transferases(HATs), which transfer acetyl groups from acetyl coenzyme A (Ac-CoA) to theε-amino group of conserved lysine residues in histone tails (Fig. 1.4). Although notas common, acetylation of lysine residues in the globular domain of histones is alsoknown to occur (Ozdemir et al., 2006). Neutralization of lysine’s positive chargedisrupts the electrostatic histone-DNA interactions, promoting the unravelingof chromatin (Kouzarides, 2007). The opposite reaction, performed by histonedeacetylases (HDACs), restores lysine’s positive charge and promotes chromatin


TCAcycle

CitrateOxaloacetateCS

Ac-CoACoA

CTP

Malate CitrateOxaloacetate

DIC

MalateMDH

Ac-CoA CoA

ACLMDH

Fatty acidsynthesis

CACT

Carnitine Acetylcarnitine

Carnitine Acetylcarnitine

Acetylcarnitine

Carnitine

Ac-CoA CoA

CAT

CoA

Ac-CoA

CAT

Histoneacetylation

NucleusCytosol

Matrix

IMM

OMM

IMS

Figure 1.5. Overview of the citrate and the carnitine shuttles. The carnitine shuttleis illustrated as being unidirectional for simplicity, but the system is fully reversible.Conversely, the citrate shuttle is an irreversible system. Abbreviations: CS, citratesynthase; CTP, citrate transport protein; DIC, dicarboxylate ion carrier; IMM, innermitochondrial membrane; IMS, intermembrane space; MDH, malate dehydrogenase;OMM, outer mitochondrial membrane. Other abbreviations are described in the text.

condensation. The overall steady-state equilibrium of histone acetylation isdetermined by the dynamic, opposing activities of histone acetyltransferases anddeacetylases.

Acetyl coenzyme A

Acetyl coenzyme A is a central metabolite found at the crossroads of multiplemetabolic pathways. It is mainly generated in the mitochondrial matrix frompyruvate and free fatty acids (FFAs), via the activities of pyruvate dehydrogenase(PDH) and the β-oxidation pathway, respectively. Peroxisomes can also generateAc-CoA via β-oxidation, but the relative contribution of this organelle in cardiactissue is believed to be insignificant (Chu et al., 1994).

Because the mitochondrial inner membrane is impermeable to Ac-CoA, theacetyl moieties must be exported to the cytosol by means of a shuttle systemin order to be used by cytosolic acetyltransferases. Acetyl groups can leavemitochondria in the form of citrate and acetylcarnitine, via the citrate andcarnitine shuttles, respectively (Fig. 1.5). In the citrate shuttle, mitochondrial

10 Introduction

citrate generated in the TCA cycle is carried to the cytosol via citrate transportprotein (CTP) and is then converted to oxaloacetate by ATP citrate lyase (ACL),generating Ac-CoA in the process. The carnitine shuttle is most commonlyimplicated in transporting long-chain FFAs into mitochondria for β-oxidation.However, it has also been proposed to act as a buffer system to prevent fluctuationsin the mitochondrial Ac-CoA/CoA-SH ratio that would otherwise affect TCAcycle dynamics (Bremer, 1983). The key proteins in the carnitine shuttle arecarnitine acetyltransferase (CAT), which interconverts Ac-CoA and acetylcarnitine,and carnitine–acylcarnitine translocase (CACT), which transports carnitine andacetylcarnitine across the mitochondrial inner membrane.

Recent studies in mammalian cells suggest that histone acetylation is linkedto mitochondrial-derived Ac-CoA via both the citrate and the carnitine shuttlesystems. Thompson and colleagues (Wellen et al., 2009) demonstrated that histoneacetylation was dependent on both glucose availability and ACL activity; simul-taneously linking the energetic status of the cell and mitochondrial metabolismto epigenetic modulation. In the same year, Madiraju et al. (2009) assessed theeffect of l-carnitine and sulfobetaine 3-08 (an inhibitor of mitochondrial carnitinetransport) supplementation in histone acetylation levels. The authors reportedthat l-carnitine supplementation increased histone acetylation levels, and thatthis effect was abolished by carnitine transport inhibition. The authors alsodemonstrated the existence of a nuclear CAT, further reinforcing the importanceof the carnitine shuttle as a donor of mitochondrial acetyl groups for nuclearacetylation reactions. Furthermore, a series of studies from the ’70s in perfused rathearts suggest that the mitochondrial and cytosolic ratios of Ac-CoA/CoA-SH andcarnitine/acetylcarnitine maintain their equilibriums under variable work loads(Oram et al., 1973; Oram et al., 1975; Idell-Wenger et al., 1978), underscoring thecarnitine-mediated link between both pools of acetyl groups. Thus, alterationsin mitochondrial Ac-CoA pools will likely reflect on the nuclear Ac-CoA pools,thereby conditioning histone acetylation dynamics.

By disrupting overall mitochondrial metabolism, DOX can be expected tointerfere with the cellular pools of acetyl groups. Indeed, it was found thatDOX treatment leads to greatly elevated Ac-CoA/CoA-SH ratios in mitochondriaisolated from rat heart (Ashour et al., 2012). This can be detrimental to cardiomy-ocyte lipid metabolism for two reasons. First, high Ac-CoA/CoA-SH ratios inhibitthe β-oxidation pathway by allosterically inhibiting 3-ketoacyl-CoA thiolase (thefinal enzyme in the β-oxidation cycle) (Schulz, 2002). Second, excess Ac-CoA willoverload the carnitine buffering system, resulting in high acetylcarnitine/carnitineratios. This limits lipid transport across the mitochondrial inner membrane, aslower levels of free carnitine are available to participate in this process.

It has recently been proposed that mitochondrial Ac-CoA levels couple cellularenergetics to nuclear gene transcription regulation (Wallace and Fan, 2010);meaning that high levels of Ac-CoA will invoke a fed state transcriptome viaepigenetic modulation. This could explain, at least in part, the overexpression ofglycolytic genes in cardiomyocytes exposed to DOX. It would also imply that cellswould have to respond to mixed signals regarding their energetic status – low ATPlevels would signal for a fasting state whereas high Ac-CoA levels would signal


for an artificial fed state. This pseudo-energy surplus signaling can be furtherexacerbated by the impairment of the energy sensor AMP-activated protein kinase(AMPK) that has been shown to follow DOX treatment (Tokarska-Schlattneret al., 2005). Clearly, the cardiac metabolic remodeling induced by DOX is amultifactorial process. It remains to be seen to which extent this effect is mediatedby aberrant histone acetylation dynamics.

1.2.2 DNA methylationDNA methylation consists in the addition of a methyl group to the 5 position ofcytosine, forming 5-methylcytosine (m5C) (Fig. 1.6). This modification occursprimarily in CpG dinucleotides, so as to ensure that methylation patterns canbe mirrored in both strands of DNA after replication. During DNA synthesis,only unmethylated cytosines are added to the growing strand. The maintenanceof the methylation pattern depends on the activity of DNA methyltransferase(DNMT) 1, which binds preferentially to hemimethylated DNA (that is, a m5CpGdinucleotide that is paired to a GpC dinucleotide)

5’-m5

CG-3’3’-GC-5’

and catalyzes the symmetrical methylation of the newly added cytosine in theopposing GpC dinucleotide (hence the CpG specificity),

5’-m5

CG-3’3’-GC

m5-5’

thus providing a way to propagate methylation patterns between cell generations(Jeltsch, 2006). Unlike DNMT1, both DNMT3a and DNMT3b methylate CpGdinucleotides with no preference for hemimethylated DNA (Okano et al., 1998).De novo methylation by the Dnmt3 family is essential for the establishment ofmammalian embryonic methylation patterns (Okano et al., 1999).

DNA regions with a high CpG density – the so-called “CpG islands” – colocalizefrequently with gene promoters or other regulatory regions required for genetranscription, and are mostly unmethylated (Guibert and Weber, 2013). Initially,DNA methylation was described as a repressive mark, since many mammaliantranscription factors bind to CpG motifs. Methylation of these motifs prevents thebinding of several transcription factors, thus repressing gene expression (Bird andWolffe, 1999). Recently, however, the outcome of DNA methylation is increasinglybeing regarded as being context-dependent. Methylation of micro RNA promoterregions is a particular example of how this epigenetic mark can lead to geneexpression activation, and has been shown to be an important mechanism inmouse embryonic development (Cui et al., 2009).

Until recently, DNA methylation was believed to be a phenomenon exclusive tonuclear DNA (nDNA). Studies performed in the last few years, however, demon-strate not only the presence of m5C in mtDNA but also a possible mitochondrial of

12 Introduction

NH

O

N

NH2

Cytosine

NH

O

N

NH2

CH3

5-Methylcytosine

SAM SAH

DNMTs

Figure 1.6. DNA methylation mechanism. The DNMT-catalyzed reaction consists inthe transfer of a methyl moiety from the universal methyl donor SAM to the 5 positionof a cytosine in a CpG dinucleotide.

DNMT1 and DNMT3a in mitochondria (Chestnut et al., 2011; Shock et al., 2011).The field of mtDNA methylation remains remarkably unexplored nonetheless.

S-Adenosyl-L-methionine

The process of DNA methylation requires the universal methyl donor S -adenosyl-l-methionine (SAM), which is synthesized through the coordinated activities of themethionine cycle and the folate cycle (Fig. 1.7). SAM is produced in the methioninecycle, where the enzyme methionine adenosyltransferase (MAT) catalyzes theaddition of methionine to the adenosyl moiety of ATP, thereby generating SAM.Removal of the methyl group in SAM by various methyltransferases producesS -adenosyl-l-homocysteine (SAH), which is hydrolyzed to homocysteine (Hcy)by SAH hydrolase (SAHH). The next step completes the methionine cycle andmerges with the folate cycle; methionine synthase (MS) tranfers the methyl groupfrom 5-methyl-tetrahydrofolate (5-MTHF) to Hcy, thus regenerating methionineand producing tetrahydrofolate (THF) as a byproduct (Iacobazzi et al., 2013).The synthesis of methionine – and SAM, by extension – is therefore dependent ona steady supply of 5-MTHF by the folate cycle.

Folate metabolism – also referred to as the one-carbon metabolism – is semi-duplicated in the cytoplasm and in mitochondria. The two compartments carryout folate metabolism for different purposes using parallel, but distinct enzymes.In the cytoplasm, folates donate one-carbon units to the synthesis of thymidylate(dTMP), purines and methionine. Mitochondrial folates are used to generateglycine, N -formylmethionyl-tRNA (fMet-tRNA) and to donate one-carbon unitsfor cytoplasmic one-carbon metabolism in the form of formate (Fox and Stover,2008). Cytoplasmic and mitochondrial folate pools do not equilibrate with eachother, but they exchange one-carbon units in the form of serine, glycine andformate (Depeint et al., 2006).

The proper folate metabolism requires some degree of coordination betweenthe two compartments. Imbalances between the two pools may arise from mito-chondrial dysfunction. For instance, mitochondrial folate metabolism is stronglyinfluenced by the NADH/NAD+ ratio. Conversion of THF to methylene-THF inmitochondria can be achieved by redox-coupled reactions, which are performed byNAD+-dependent dehydrogenases, and by a non-redox-coupled reaction (Fig. 1.7).


Cytoplasm

Mitochondria

Methioninecycle

SAMMAT

MT

SAHH*

MS

Met

HcySAH

PPi + PiATP

Folatecycle

THF

MTFR

cSHMT

Methylene-THF

DHF

dUMP

dTMPTS

5-MTHF

sergly

THF

GCS*

THF

mSHMT

MTF

SarDH* DMGDH*

Methylene-THF

MTHFD*

Methenyl-THF

MTHFC

Formyl-THFFTHFS

THF

FormatefMet-tRNA

gly ser

Cystathionine

Methylationreactions

Formate

THF Formyl-THF

DHF

Folate

Glutathionesynthesis

Purinesynthesis

ser

CBS

DHFR

DHFR

C1-THFS

Figure 1.7. Simplified overview of folate-mediated one-carbon metabolism. Theconnection between methionine metabolism and glutathione biosynthesis is also shown.Asterisks denote NAD(P)+-dependent enzymes. Abbreviations: C1-THFS, C1-THFsynthase; CBS, cystathionine β-synthase; DHF, dihydrofolate; DHFR, DHF reductase;DMGDH, dimethylglycine dehydrogenase; FTHFS, formyltetrahydrofolate synthetase;GCS, glycine cleavage system; MFT, methionyl-tRNAMet

f formyltransferase; MT, gener-alized methyltransferase; MTFR, methylene-THF reductase; MTHFC, methenyl-THFcyclohydrolase; MTHFD, methylene-THF dehydrogenase; SarDH, sarcosine dehydroge-nase; TS, thymidylate synthase. Other abbreviations are described in the text.

Impairment of the ETC decreases the mitochondrial oxidation of NADH to NAD+,thus increasing the NADH/NAD+ ratio. This limits the entry of THF into themitochondrial folate cycle via redox-coupled reactions, forcing mitochondria to relyprimarily on one-carbon units from serine via the activity of mitochondrial serinehydroxymethyl transferase (mSHMT). As a result, cytosolic serine is transfered tomitochondria to support mitochondrial folate metabolism, slowing down in turncytoplasmic folate metabolism at the cytosolic serine hydroxymethyl transferase(cSHMT) step. This has a propagating effect in the methionine cycle by hinderingthe cytoplasmic folate cycle, thus decreasing the pools of 5-MTHF that feed the

14 Introduction

synthesis of methionine (Naviaux, 2008). Evidence in favor of this reasoning comesfrom an in vitro study using ρ0 cells, which are devoid of mtDNA (Smiragliaet al., 2008). Depletion of mtDNA compromises OxPhos metabolism since mtDNAencodes for 11 subunits of the ETC complexes (I, III and IV) and 2 subunits ofATP synthase (Taanman, 1999). The authors reported significant alterations inthe methylation patterns of several genes, which were only partially reversed uponre-introduction of wild-type mitochondria.

An alternative route to divert substrates away from the methionine cycle isthe glutathione (GSH) biosynthetic pathway. De novo synthesis of GSH requirescysteine, which can be derived from Hcy and serine in the transsulfuration pathway(Lu, 2009). Depletion of GSH pools by excessive oxidative stress increases the fluxof Hcy to the GSH biosynthetic pathway, thus diverting Hcy and serine away fromthe methionine salvage pathway and the cytoplasmic folate cycle, respectively.

Doxorubicin can be expected to interfere with SAM metabolism by bothmechanisms discussed above. Increases in the NADH/NAD+ ratio would arisefrom ETC impairment due to protein carbonylation and mtDNA oxidation. Tworecent studies, one using an in vitro model (Wang et al., 2012) and the other aperfused heart system (Montaigne et al., 2010), provide indirect evidence thatDOX increases the NADH/NAD+ ratio in cardiomyocytes. Depletion of GSHwould result from excessive oxidative stress; such has been shown to occur incardiac tissue of rats treated with DOX (Zhou et al., 2001b; Sadik et al., 2008).Interestingly, a study from the early ’90s reported a significant protective effect inDOX-treated rats that were co-administered SAM (Russo et al., 1994). However,whether this effect is due to epigenetic maintenance, increased GSH levels or someother factor remains to be investigated.

1.3 ObjectivesThe molecular connection between cellular metabolism and epigenetic modifi-cations has been discussed above and is summarized in Fig. 1.8. Given thatDOX disrupts normal cardiomyocyte metabolism, the overall hypothesis behindthis project is that DOX disturbs the pool of metabolites directly or indirectlyinvolved in epigenetic modulation, thereby altering normal cardiomyocyte epi-genetic patterns in a non-reversible manner upon withdrawal of the drug andultimately locking the cardiomyocyte in a pathological transcriptional phenotype.This would account for many observations described in the literature and indeedfor the persistent nature of DOX-induced cardiotoxicity.

Having in mind the background described and our general hypothesis, theobjective of the present work is to verify if there is a correlation between mito-chondrial dysfunction, epigenetic disruption and alteration to the transcriptome– all of which are necessary to be in place to corroborate our general and morefar-reaching hypothesis.

1.3 Objectives 15

TCA cycle

CitrateGlucoseGlycolysis

H1

OAADPrSIRTs

NAD+

Ac-CoA

HATs

DNMTs

SAM

Methionine

NAD+ NADH

Folatemetabolism

Ac-CoA FFA

β-oxidation

Acetyl lysine

5-Methylcytosine

Cytosine

Figure 1.8. Relationship between cellular metabolism and epigenetic modifications.The glycolytic flux affects the NADH/NAD+ ratio, which controls the activity ofsirtuin histone deacetylases. Mitochondrial citrate and Ac-CoA levels determine thelevels of cytosolic Ac-CoA that is used as an acetyl donor for HAT-mediated histoneacetylation. Finally, the synthesis of SAM, the methyl donor for DNMT-mediatedDNA methylation, is dependent on the metabolism of methionine, which in turn isindirectly regulated by mitochondrial folate metabolism. Abbreviations: H1, histoneH1; OAADPr, O-acetyl-ADP ribose. Other abbreviations are described in the text.

Chapter 2

Materials and Methods

2.1 Animals and treatment protocolMale Wistar Han rats were purchased form Charles River Laboratories (France)with 7 weeks of age. One week prior to the initiation of the experiment, animalswere acclimated and maintained in local animal house facilities (CNC – Faculty ofMedicine, University of Coimbra, Coimbra, Portugal). Animals were group-housedin type III-H cages (Tecniplast, Italy) with irradiated corn cob grit bedding (ScobisDue, Mucedola, Italy) and environmental enrichment, maintained in controlledenvironmental requirements (22 ℃, 45–60 % humidity, 15–20 air changes/hour,12 hours artificial light/dark cycle, noise level < 55 dB) and free access to standardrodent food (4RF21 GLP certificate, Mucedola, Italy) and HCl-acidified water(pH 2.6).

Experimental manipulation was initiated with 8 weeks old rats weighing220–260 g. Animals were randomly assigned to two experimental groups: saline(SAL)-treated (n = 6) and DOX-treated (n = 6). Rats received seven weeklysubcutaneous injections of DOX (2 mg kg−1), dissolved in normal saline (0.9 %[w/v] NaCl), or an equivalent volume of SAL solution. All animals were injectedand weighed during the light phase of the cycle. Animals were euthanized bycervical dislocation followed by decapitation two weeks after the last injection(Fig. 2.1). ∗

2.2 Subcellular fractionationSubcellular fractions for Western blotting analyses were obtained from heart andliver tissue according to the method described by Dimauro et al. (2012), with slightmodifications. The procedure is summarized in Fig. 2.2. Excised tissue was weighed(20 mg), rinsed with PBS, resuspended in ice-cold sucrose-Tris-magnesium (STM)buffer (50 mM Tris-HCl [pH 7.4], 250 mM sucrose, 50 mM MgCl2; supplementedfreshly with 1 mM dithiothreitol [DTT], 1µL mL−1 protease inhibitor cocktail

∗The animal treatment protocol and animal data shown in Fig. 3.1 and Table 3.1 wasperformed by Filipa S. Carvalho (FELASA category B accredited), Ana Burgeiro and RitaGarcia.

17

18 Materials and Methods

8 9 10 11 12 13 14 15 16Age:(weeks)

n = 6

SALO O O O O O O �

♂ Wistar Han rats

n = 6

DOXH H H H H H H �

Figure 2.1. Timeline of the treatment protocol. Animals were given seven weeklysubcutaneous injections of DOX (2 mg kg−1; ▼) or of SAL solution (▽). All animalswere euthanized at 16 weeks of age (�).

[PIC; Sigma–Aldrich, Barcelona, Spain; #P8340], 5 mM nicotinamide [NAM] and5 mM sodium butyrate [SB]) and homogenized on ice in a pre-cooled Potter–Elvehjem glass homogenizer using a tight-fitting Teflon pestle (wall clearance:0.10 mm) attached to a mechanical overhead stirrer (Heidolph, Germany) set to600 rpm for 50–60 strokes. The homogenate was decanted into a microcentrifugetube and maintained on ice for 15 min, vortexed at maximum speed for 15 s andcentrifuged at 800 g for 15 min at 4 ℃. The supernatant was labeled as S1 andused for concurrent extraction of cytosolic and mitochondrial fractions. The pelletwas washed thrice with centrifugation steps after resuspension in STM buffer.The washed pellet was then resuspended in NaCl-EDTA-HEPES (NEH) buffer(0.5 M NaCl, 0.2 mM EDTA, 20 mM HEPES, pH 7.9, 1.5 mM MgCl2, 20 % [v/v]glycerol, 1 % [v/v] Triton X-100; supplemented as before) , vortexed at maximumspeed for 15 s and maintained on ice for 30 min. Nuclei were lysed with sonication(3× in 10 s bursts) in a water bath sonicator with 30 s pauses between sonicationsteps whilst keeping on ice throughout. The lysate was centrifuged for 30 min at9,000 g and the resulting supernatant (the nuclear fraction) was stored at −80 ℃until use.

Concurrently, cytosolic and mitochondrial fractions were obtained from thesupernatant S1 by centrifugation at 800 g for 10 min, so as to remove any remainingnuclear contamination. If the nuclear pellet P2 was substantial, it was resuspendedin STM buffer and combined with the pellet P1.1. The supernatant S2 containingthe cytosolic and mitochondrial fractions was centrifuged for 10 min at 11,000 g, thesupernatant S3 was precipitated in cold 80 % acetone supplemented with 1 M NaClat −20 ℃ for 1 h (Crowell et al., 2013) and subsequently centrifuged for 10 min at11,000 g. The resulting pellet P4 (the cytosolic fraction) was resuspended in STMbuffer and stored at −80 ℃ until use. The mitochondrial pellet P3.1 was washedonce with a centrifugation step after resuspension in STM buffer, the washedpellet was then resuspended in Tris-EDTA with Triton X-100 (TE-TX-100) buffer(50 mM Tris-HCl [pH 6.8], 1 mM EDTA, 0.5 % [v/v] Triton X-100; supplementedas before), sonicated as before and stored at −80 ℃ until use.

Nuclear fractions for enzymatic activity assays were obtained from heart and

2.3 Subcellular fractionation 19

·Homogenize 50–60 strokesat 600 rpm in STM buffer·Decant homogenate to a

microcentrifuge tube

·Incubate on ice for 30′

·Vortex 15′′ max speed·Centrifuge for 15′ at 800 g

P1.1

·Resuspend in STM buffer·Vortex 15′′ max speed

·Centrifuge for 15′ at 500 g

·Centrifuge for 10′ at 800 g

S1

P2

·Centrifuge for 10′ at 11,000 g

S2

·Combine with P1.1

P1.2



Discard

P3.1



·Precipitate in 1M NaCl80% acetone·Centrifuge for 10′ at 11,000 g

S3

P3.2

·Resuspend in TE-TX-100 buffer·Sonicate 3× 10′′

Discard

Mitochondrial fraction

P4

·Resuspend in STM buffer

Discard

Cytosolic fractionP1.3

·Resuspend in NEH buffer·Vortex 15′′ max speed·Incubate on ice for 30′

·Sonicate 3× 10′′


Discard

P1.4

Discard

Nuclear fraction

Figure 2.2. Schematic workflow of the subcellular fractionation protocol. The dashedarrow denotes an optional step.

liver tissue as described above, but with the omission of buffer supplementation.Protein content of all extracts was determined by the BCA assay, calibrated withbovine serum albumin.


2.3 DNA extraction

Total genomic DNA for mtDNA and m5C quantification was extracted fromheart and liver tissue using the PureLink™ Genomic DNA Mini Kit (Invitrogen),according to the manufacturer’s instructions. Briefly, excised tissue was weighed(15–20 mg) and incubated with digestion buffer and proteinase K (NZYTech,Portugal; MB01902). The lysate was centrifuged and the resulting supernatant wastreated with RNase A. After incubation, the lysate was mixed with binding bufferand ethanol and centrifuged through a silica-based spin column. After washingthe column, bound DNA was eluted by centrifuging elution buffer through thecolumn. The extracted DNA samples were quantified spectrophotometrically atA260 nm using a NanoDrop 2000 spectrophotometer (Thermo Scientific, USA) andstored at −80 ℃ until use.

2.4 RNA extraction and cDNA synthesis

Total RNA was extracted from heart and liver tissue using the Aurum™ TotalRNA Mini Kit (Bio-Rad, Hercules, CA, USA), according to the manufacturer’sspecifications. Briefly, excised tissue was weighed (15–20 mg) and immediatelyhomogenized in lysis solution using a T 25 Ultra-Turrax® dispersion device (IKA®,Germany). After centrifuging the lysate at 16,000 g for 3 min, the resulting super-natant was diluted 1:1 with 60 % ethanol and centrifuged through a silica-basedspin column at 16,000 g for 1 min. Following a washing step, the column wasincubated with DNase I for 25 min at room temperature. Additional washingsteps were performed before the column-bound RNA was eluted by centrifugingthe provided elution solution through the column. Thereafter, RNA qualityand integrity was ascertained by visualization of the 18 S/28 S rRNA band pat-tern using an Experion™ Automated Electrophoresis System (Bio-Rad). TotalRNA was quantified spectrophotometrically at A260 nm using a NanoDrop 2000spectrophotometer (Thermo Scientific) and stored at −80 ℃ until use.

First-strand complementary DNA (cDNA) synthesis was performed using theiScript™ cDNA synthesis kit (Bio-Rad), according to the manufacturer’s specifica-tions. Briefly, oligo(dT)/random hexamer primer mediated reverse transcriptionwas carried out in reactions comprising 1µg of total RNA, 1× iScript™ reactionmix and iScript™ reverse transcriptase, made up to 20µL with nuclease-free water,under the following cycling parameters:

25℃5:00

42℃30:00

85℃5:00

2.8 HAT activity assay 21

2.5 HAT activity assayHistone acetyltransferase activity in cardiac and liver tissue was measured usingthe colorimetric Histone Acetyltransferase Assay Kit (Abcam; ab65352), accordingto the manufacturers’ instructions. This kit measures the acetylation rate of apeptide substrate by active HATs. This process releases free CoA, which thenserves as a coenzyme for the production of NADH by a NADH-generating enzyme(undisclosed). The rate of NADH production is measured spectrophotometricallythrough the reduction of a tetrazolium salt into a colored formazan product.Briefly, 25µg of nuclear extract were incubated with HAT substrates I and IIand NADH-generating enzyme in assay buffer for 2 h at 37 ℃. Absorbance wasmeasured at 440 nm using a Cytation™ 3 microplate reader (BioTek, USA).

2.6 HDAC activity assayHistone deacetylase activity in cardiac tissue was measured using the fluorimet-ric Histone Deacetylase Activity Assay Kit (Abcam; ab156064), according tothe manufacturers’ instructions. This kit measures the deacetylation rate of afluorescence-labeled acetylated peptide substrate by active HDACs.

X−X−X−(Ac)K−MCAHDACs−−−−→H2O

X−X−X−K−MCA + AcOH {2.1}

Upon deacetylation, the substrate peptide is cleaved by a peptidase (lysyl en-dopeptidase, LE)

X−X−X−K−MCALE−−→ X−X−X−K + AMC {2.2}

and the fluorescent group (7-amino-4-methyl coumarin, AMC) is released. Themeasured fluorescence is directly proportional to the activity of HDACs. The speci-ficity of the assay was ascertained with negative controls containing trichostatin A– a selective inhibitor of class I and II HDACs. Briefly, 25µg of nuclear extract wereincubated with the substrate peptide and endopeptidase in assay buffer for 30 minat room temperature. Fluorescence was measured at λEx/λEm = 355/460 nmusing a Cytation™ 3 microplate reader (BioTek, USA).

2.7 Enzyme-linked immunosorbent assay (ELISA)of global DNA m5C content

The m5C content of total DNA from heart and liver tissue was determined using thefluorometric EpiSeeker Methylated DNA Quantification Kit (Abcam; ab117129),according to the manufacturer’s instructions. Briefly, 100 ng of DNA were boundto the provided assay wells, incubated for 60 min with capture antibody and thenfor 30 min with detection antibody. After incubating the probed DNA with theprovided substrate for 4 min, methylated DNA was quantified fluorimetrically byreading the fluorescence at λEx/λEm = 544/590 nm in a Victor™ X3 multilabelplate reader (PerkinElmer). Results were expressed as percent m5C, calculatedusing a linear regression equation generated with controls provided in the kit.


2.8 Primer designPrimers used in quantitative real-time PCR for nDNA-encoded gene transcriptswere designed using the Primer-BLAST online tool (NCBI) with the followingspecifications: PCR product size 90–200 bp; exon-exon junction span enforcedwhen possible. For mtDNA genes and respective transcripts, primers were designedusing the Beacon Designer™ 8.0 software (Premier Biosoft International). For thenDNA β2M (B2m) gene, primers were designed using Primer-BLAST with thefollowing specifications: PCR product size 90–200 bp; exon-intron junction spanenforced. All primer sequences used in this study are shown in Table 2.1.

2.9 Quantitative real-time PCRQuantitative real-time PCR (qPCR) was performed in duplicate reactions, eachcomprising 12.5 ng of either cDNA or DNA, 500 nM of each gene-specific primerand 1× SsoFast™ EvaGreen® Supermix (Bio-Rad), made up to 10µL with nuclease-free water. Reactions were performed in sealed 96-well plates and monitored witha CFX96™ Real-Time PCR Detection System (Bio-Rad). Results were analysedusing the CFX Manager™ 3.1 software (Bio-Rad). For gene expression analyses,reactions were performed with the following cycling parameters:

95℃0:30

95℃0:05

60–63℃0:05

40 cycles

To confirm the absence of genomic DNA contamination, a negative control wasincluded in the reverse transcriptase reaction. Genomic DNA contamination wasconsidered insignificant when the cycle threshold difference between the no-reversetranscriptase and the respective sample was above 10 cycles. Transcript levels werenormalized relative to the expression of three housekeeping genes (18 S [Rn18s],TBP [Tbp] and β-actin [Actb]) using the Pfaffl method (Pfaffl, 2001). For mtDNAcopy number quantification, reactions were performed with the following cyclingparameters:

98℃2:00

98℃0:05

63℃0:05

40 cycles

Levels of the mitochondrial COX1 (Mt-co1 ) gene were normalized relative to thelevels of the single-copy nuclear β2M gene, also using the Pfaffl method. For allreactions, amplification of a single PCR product was confirmed by melt curve

2.9Q

uantitativereal-tim

eP

CR

23

Table 2.1List of oligonucleotide primer sequences used for qPCR. Symbols and names are relative to the Rat Genome Database (RGD,http://rgd.mcw.edu/); accession numbers are relative to the GenBank® database (Benson et al., 2014). For each primer, F and R indicateforward and reverse orientation, respectively.

Target Symbol Name Accession no. Sequence

18 S Rn18s 18S ribosomal RNA NR_046237 F 5’-ACTCAACACGGGAAACCTC-3’R 5’-ACCAGACAAATCGCTCCAC-3’

ACAA2 Acaa2 acetyl-CoA acyltransferase 2 NM_130433 F 5’-CCTCAGTTCTTGGCTGTTCA-3’R 5’-CCACCTCGACGCCTTAAC-3’

ACC2 Acacb acetyl-CoA carboxylase beta NM_053922 F 5’-ACAGCAAGAGCCTTCAAGAG-3’R 5’-CGCAGTCCGTAATCCACAA-3’

ACL Acly ATP citrate lyase NM_016987a F 5’-GCTGCGTTACCAAGACACT-3’R 5’-AACTGGACCTCAGAAGAGAACA-3’

ACOX1 Acox1 acyl-CoA oxidase 1, palmitoyl NM_017340 F 5’-GCAGACAGCCAGGTTCTTGATG-3’R 5’-ACTCGGCAGGTCATTCAGGTAT-3’

ATP8 Mt-atp8 mitochondrially encoded ATP synthase 8 AC_000022b F 5’-TGCCACAACTAGACACATCCACAT-3’R 5’-GAGGGAGGTGCAGGAAAGGTT-3’

CACT Slc25a20 solute carrier family 25 (carnitine/acylcarnitinetranslocase), member 20

NM_053965 F 5’-GGAGAACGGATCAAATGCTTACT-3’R 5’-GGCAGGAACATCTCGCAT-3’

CBP Crebbp CREB binding protein NM_133381 F 5’-GCACCCATGCCAACAACATT-3’R 5’-ACCTGGCCCTGTGAAACAC-3’

COX1 Mt-co1 mitochondrially encoded cytochrome c oxidase I AC_000022b F 5’-GCCAGTATTAGCAGCAGGTATCA-3’R 5’-GCCGAAGAATCAGAATAGGTGTTG-3’

Cyt b Mt-cyb mitochondrially encoded cytochrome b AC_000022b F 5’-TACGCTATTCTACGCTCCATTC-3’R 5’-GCCTCCGATTCATGTTAAGACTA-3’

Cyt c Cycs cytochrome c, somatic NM_012839 F 5’-CTTGGGCTAGAGAGCGGGAC-3’R 5’-CCCATTTTAAATTCGGTCCGGG-3’

DNMT1 Dnmt1 DNA (cytosine-5-)-methyltransferase 1 NM_053354 F 5’-GGAAGGTGAGCATCGACGAA-3’R 5’-TGCTGTGACCCTGGCTAGAT-3’

Ep300 Ep300 E1A binding protein p300 XM_001076610 F 5’-TGGAGGCACTTTACCGACAG-3’R 5’-GATCCATGGGGCTCTTCACA-3’

aAmplifies other transcript variants. (continued on next page)bReference sequence for the complete Wistar strain mitochondrial genome.

http://rgd.mcw.edu/

http://www.ncbi.nlm.nih.gov/nuccore/NR_046237

http://www.ncbi.nlm.nih.gov/nuccore/NM_130433




http://www.ncbi.nlm.nih.gov/nuccore/AC_000022







http://www.ncbi.nlm.nih.gov/nuccore/XM_001076610

24M

aterialsand

Methods

Table 2.1 (continued)


HDAC1 Hdac1 histone deacetylase 1 NM_001025409 F 5’-AGCCAAAGGGGTCAAAGAAGA-3’R 5’-TGAGAAATTGAGGGAAAGTAAGGGA-3’

HDAC2 Hdac2 histone deacetylase 2 NM_053447 F 5’-GGAGGTCGTAGGAATGTCGC-3’R 5’-TTTGGCTCCTTTGGTGTCTGT-3’

HDAC3 Hdac3 histone deacetylase 3 NM_053448 F 5’-CTGGGAGGTGGTGGTTACAC-3’R 5’-TCTGATTCTCGATGCGGGTG-3’

HDAC4 Hdac4 histone deacetylase 4 NM_053449 F 5’-CATGGTTCAACGTTCATCTCTGC-3’R 5’-CGCAAACTCGAAGTCCCACT-3’

HDAC5 Hdac5 histone deacetylase 5 NM_053450 F 5’-CCCGTCCGTCTGTCTGTTAT-3’R 5’-CCTGACATGCCATCCGACTC-3’

HDAC6 Hdac6 histone deacetylase 6 XM_001057931 F 5’-CCCTACAGGAGGTGGAGTTG-3’R 5’-GCTCCTGGAACGGCTCTTT-3’

HDAC7 Hdac7 histone deacetylase 7 XM_001059057 F 5’-GACAAGAGCAAGCGAAGTGC-3’R 5’-ACTGTTCTCTCAAGGGCTGC-3’

HDAC8 Hdac8 histone deacetylase 8 NM_001126373 F 5’-GTGCTGGAAATCACGCCAAG-3’R 5’-CACATGCTTCAGATTCCCTTTGA-3’

HDAC9 Hdac9 histone deacetylase 9 NM_001200045 F 5’-AAACCCAGCATCTGACCTCCA-3’R 5’-CTGATTTCACGTCCACTGAGCTG-3’

HDAC10 Hdac10 histone deacetylase 10 NM_001035000 F 5’-GCCCATAGAGGTCAGAGGAGA-3’R 5’-AGGCTAAGGGCAGTACCAGA-3’

MAT2a Mat2a methionine adenosyltransferase II, alpha NM_134351 F 5’-GGGGAAGGTCATCCAGATAAGA-3’R 5’-AGCAACAGTTTCACAAGCCAC-3’

ND1 Mt-nd1 mitochondrially encoded NADH dehydrogenase 1 AC_000022a F 5’-TCCTCCTAATAAGCGGCTCCTTCT-3’R 5’-TGGTCCTGCGGCGTATTCG-3’

ND6 Mt-nd6 mitochondrially encoded NADH dehydrogenase 6 AC_000022a F 5’-TCCTCAGTAGCCATAGCAGTTGT-3’R 5’-GTTGTCTAGGGTTGGCGTTGAA-3’

NDUFA9 Ndufa9 NADH dehydrogenase (ubiquinone) 1 alphasubcomplex, 9

NM_001100752 F 5’-GAGAAGCAAGGCAGTGGGAGAG-3’R 5’-ACGAGAGGCACAGCAAGAAACC-3’

aReference sequence for the complete Wistar strain mitochondrial genome. (continued on next page)















2.9Q

uantitativereal-tim

eP

CR

25

Table 2.1 (continued)


NRF-1 Nrf1 nuclear respiratory factor 1 NM_001100708 F 5’-CCAAGCATTACGGACCATAGTT-3’R 5’-CAGTACCAACCTGGATGAGC-3’

NRF-2α Gabpa GA binding protein transcription factor, alphasubunit

NM_001108841 F 5’-CGGCACCAAGCATATCAC-3’R 5’-GGACCACTGTATAGGATCATAGG-3’

NRF-2β Gabpb1 GA binding protein transcription factor, beta subunit1

NM_001039036 F 5’-GCACCGCTGTCATTTTGTCG-3’R 5’-CAGAGCAGTCTCACGTCCTC-3’

PCAF Pcaf p300/CBP-associated factor XM_003750617 F 5’-CAGGTCAAGGGCTATGGAACC-3’R 5’-CCCATCAAAGTGGCTCCTTCA-3’

PGC-1α Ppargc1a peroxisome proliferator-activated receptor gamma,coactivator 1 alpha

NM_031347 F 5’-TGAGAAGCGGGAGTCTGAAAG-3’R 5’-AACCATAGCTGTCTCCATCATCC-3’

POLγ Polg polymerase (DNA directed), gamma NM_053528 F 5’-ATAATGGCCCACACCCGTTT-3’R 5’-GAGTTTCCGGTACCACCCAG-3’

PPARα Ppara peroxisome proliferator activated receptor alpha NM_013196 F 5’-AGACTAGCAACAATCCGCCTTT-3’R 5’-TGGCAGCAGTGGAAGAATCG-3’

SDHA Sdha succinate dehydrogenase complex, subunit A,flavoprotein (Fp)

NM_130428 F 5’-CTATGGAGACCTACAGCATCT-3’R 5’-AATCCGCACCTTGTAATCTTC-3’

TAZ Taz tafazzin NM_001025748 F 5’-CAAATGGGGAATTGGACGGC-3’R 5’-AGGGAGTGTACTGAAGGGCT-3’

TBP Tbp TATA box binding protein NM_001004198 F 5’-CCTATCACTCCTGCCACACC-3’R 5’-CAGCAAACCGCTTGGGATTA-3’

TFAM Tfam transcription factor A, mitochondrial NM_031326 F 5’-AATGTGGGGCGTGCTAAGAA-3’R 5’-TCGGAATACAGATAAGGCTGACAG-3’

UQCRC1 Uqcrc1 ubiquinol-cytochrome c reductase core protein I NM_001004250 F 5’-GAGACACAGGTCAGCGTATTG-3’R 5’-TTTGTTCCCTTGAAAGCCAGAT-3’

UQCRFS1 Uqcrfs1 ubiquinol-cytochrome c reductase, Rieske iron-sulfurpolypeptide 1

NM_001008888 F 5’-GGACGTGAAGCGACCCTT-3’R 5’-CGAACAGAAGCAGGAACATTCAG-3’

β2M B2m beta-2 microglobulin 24223a F 5’-AGAGAACTCAACGGTGGCA-3’R 5’-CGACCGCACACTATAGGGAC-3’

β-actin Actb actin, beta NM_031144 F 5’-AGATCAAGATCATTGCTCCTCCT-3’R 5’-ACGCAGCTCAGTAACAGTCC-3’

aGene ID.














http://www.ncbi.nlm.nih.gov/gene/24223



analyses and PCR product 2 % agarose gel electrophoresis at the end of everyexperiment. ∗

2.10 Mitochondrial DNA integritymtDNA integrity was evaluated with a PCR-based assay adapted from Kovalenkoand Santos (2009) and Bliksoen et al. (2015). The rationale for this assay isbased on the ability of some lesions (e.g. single- and double-strand breaks) toblock the progression of the polymerase on DNA templates, resulting in decreasedamplification of the target amplicon. Assuming a Poisson distribution, whichmodels that DNA lesions are randomly distributed, amplification of long templatesis selectively inhibited, since short templates have a lower probability of havinglesions and are therefore considered as representative of undamaged DNA. Becauseonly a single lesion per strand is necessary to block the polymerase, the ratiobetween a long fragment and a short fragment effectively measures the fraction ofundamaged template molecules.

Two separate PCR reactions were performed – one amplifying a long 10,821 bp(10 kb) fragment of mtDNA and the other amplifying a short 121 bp fragment ofmtDNA. For the 10 kb fragment, primer sequences used were the forward andreverse primers of the ND1 (Mt-nd1 ) and ND6 (Mt-nd6 ) targets, respectively(see Table 2.1). PCR was performed in reactions comprising 50 ng of total DNA,500 nM of each primer and 1× Phusion™ Hot Start II High-Fidelity PCR MasterMix (Thermo Scientific), made up to 25µL with nuclease-free water, under thefollowing cycling parameters:

98℃2:00

98℃0:10

68℃0:20

72℃5:45

72℃7:30

18 cycles

For the 121 bp fragment, primer sequences used were the forward and reverseprimers of the COX1 target (see Table 2.1). PCR was performed in reactionscomprising 50 ng of total DNA, 500 nM of each primer and 1× HotStarTaq®

Master Mix (Qiagen, Germany), made up to 25µL with nuclease-free water, underthe following cycling parameters:

95℃2:00

94℃0:10

63℃0:30

72℃1:00

72℃10:00

18 cycles

∗Andreia Vilaça assisted with qPCR experiments.

2.12 Western blotting 27

For both fragments, a control reaction containing 50 % (25 ng) of total DNA wasincluded to confirm that the PCR was terminated in the exponential phase. Afterdiluting the 10 kb and 121 bp PCR mixtures in loading buffer (2.5 % [w/v] Ficoll400), equivalent volumes of the diluted mixtures (5µL) alongside 1 kb and 100 bpDNA ladders were loaded onto 0.7 % and 2 % agarose gels stained with GreenSafePremium (NZYTech; MB13201) and separated electrophoretically at 100 V for30 min. After the run, gels were imaged using a Benchtop UV Transilluminator(UVP, LLC Upland, CA, USA; Cambridge, UK) and a BioSpectrum® 500 imagingsystem (UVP). PCR products were quantified by densitometric scanning asdetailed below. mtDNA integrity was determined by calculating the ratio betweenthe 10 kb fragment and the 121 bp fragment.

2.11 Western blottingSubcellular fractions from ventricular heart tissue, obtained as described above,were denatured in Laemmli sample buffer (60 mM Tris-HCl [pH 6.8], 2 % [w/v]SDS, 10 % [v/v] glycerol, 100 mM DTT, 0.01 % [w/v] bromophenol blue) at95 ℃ for 5 min. Equivalent amounts of protein (15µg) alongside a pre-stainedprotein marker of known molecular weight (NZYTech; MB09002) were loadedonto 10 %, 1.5 mm handcast SDS-PAGE gels and separated electrophoreticallyat a constant voltage of 120 V until the bromophenol blue dye front reached thebottom of the gel. Proteins were electrotransfered from the electrophoresis gelonto polyvinylidene difluoride (PVDF) membranes (Millipore, USA) at 100 Vfor 90 min at 4 ℃. After transfer, the PVDF membranes were briefly rinsed indistilled water and incubated in Ponceau S staining solution (0.1 % [w/v] in 5 %[v/v] acetic acid) for 1 min, followed by destaining in distilled water to removenon-specific Ponceau S staining. The membranes were then inserted in betweentransparency sheets and imaged using a BioSpectrum® 500 imaging system (UVP).After destaining in Tris-buffered saline with Tween (TBS-T: 10 mM Tris-HCl[pH 7.4], 150 mM NaCl and 0.05 % [v/v] Tween 20), membranes were blocked with5 % fat-free milk in TBS-T for 1 h at room temperature and then incubated withacetyl-lysine (Ac-Lys) antibody (1:1000; Cell Signaling Technology; #9441) at 4 ℃overnight. Membranes were then washed with TBS-T and incubated for 2 h at 4 ℃with alkaline phosphatase (AP)-conjugated goat anti-rabbit IgG (1:5000; SantaCruz Biotechnology; sc-2007). After a final wash in TBS-T, immunoreactiveproteins were detected by membrane exposure to the enhanced chemifluorescence(ECF) reagent (GE Healthcare, Piscataway, NJ, USA) followed by imaging witha BioSpectrum® 500 imaging system (UVP). Quantification by densitometricscanning (see below) was normalized to protein levels determined by Ponceau Sstaining (Romero-Calvo et al., 2010).

2.12 Densitometric analysisPCR product electrophoresis, Western blot and Ponceau S staining images werequantified by densitometric scanning in ImageJ v. 1.49a (Schindelin et al., 2012).


For PCR product electrophoresis and Western blots, gels and membranes wereimaged and saved as 8-bit grayscale TIFF files. Selected bands were adjustedfor background noise manually using the straight line selection tool and theirintensities were quantified using the gel analyzer tool. For Ponceau S staining,membranes were imaged and saved as before and adjusted for background noiseusing the rolling ball algorithm. Lanes were quantified using the gel analyzertool. The linearity of Western blot and Ponceau S measurements was confirmedby including a control well with 50 % of the loaded protein – bands, lanes andregions displaying measurements within 40 % to 60 % of the respective undilutedsample were considered quantifiable. For Ac-Lys quantification, the densities ofall quantifiable immunoreactive protein bands were merged.

2.13 Statistical analysisAll data are expressed as means ± SEM. The normality of distributions and thehomogeneity of variances were objectively evaluated by the Shapiro–Wilk testand the F -test, respectively. Comparisons between two groups were performedusing the Student t-test, the Welch t-test, the Mann–Whitney U test or Yuen’stest for trimmed means, as appropriate (Wilcox, 2012). A two-sided p < 0.05 wasconsidered statistically significant. All statistical analyses were performed usingthe R v. 3.1.2 statistical software environment (R Core Team, 2014).

Chapter 3

Results

3.1 Body, heart and liver weightRats were administered seven weekly injections of saline solution or DOX (2 mg kg−1)and sacrificed two weeks following the last injection. All but one animal survivedthe entire treatment protocol. DOX-treated rats weighed significantly less thancontrol rats at sacrifice (14 %; Table 3.1) and throughout most of the duration ofthe study (Fig. 3.1). No significant changes in either heart or liver weight wereobserved between the two treatment groups (Table 3.1). Therefore, the increasein organ:body ratios seen in DOX-treated animals reflects a change in body massrather than alterations in organ mass (Table 3.1).

3.2 DOX alters mitochondrial biogenesis tran-scriptomics and mitoepigenomics

Subchronic DOX treatment alters cardiac mitochondrial function (Pereira et al.,2012). To assess if alterations to the mitochondrial biogenesis transcriptome couldplay a role in mitochondrial alteration, we measured transcripts of a numberof genes involved in this process, including peroxisome proliferator-activated(PPAR) receptor gamma coactivator 1-alpha (PGC-1α), the master regulator of

Table 3.1Impact of DOX treatment on body, heart and liver weights in rats

Body weight(g)

Heart weight(g)

Liver weight(g)

Heart:bodyratio (×103)

Liver:bodyratio (×103)

SAL 408.0 ± 18.09 1.35 ± 0.09 13.0 ± 0.7 3.29 ± 0.17 32.0 ± 0.70DOX 351.7 ± 6.00∗ 1.43 ± 0.04 13.7 ± 0.6 4.07 ± 0.11∗∗ 39.0 ± 1.65∗∗

Heart, liver and body weights of rats treated with seven weekly injections of 2 mg kg−1 DOX oran equivalent volume of saline. Data are presented as means ± SEM of values from 5–6 animalsin each experimental group.

∗p < 0.05.∗∗p < 0.01.

29

30 Results

1 2 3 4 5 6 7 8 9200

300

400

500

∗∗∗∗

∗

Weeks

Bod

ym

ass

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SAL DOX

Figure 3.1. Effect of DOX treatmenton body mass gain. Lines represent themeans±SEM of each experimental group ateach time point. ∗p < 0.05 vs. correspond-ing time point.

mitochondrial biogenesis, and its main downstream transcription factors (nuclearrespiratory factors [NRF] -1 and -2) (Scarpulla, 2011). Subchronic DOX treatmentdecreased the transcript levels for PGC-1α (Ppargc1a), but did not affect NRF-1(Nrf1 ) and the alpha subunit of NRF-2 (NRF-2α [Gabpa]) mRNAs. In contrast,NRF-2β (Gabpb1 ) mRNA was significantly increased (Fig. 3.2A). Because mtDNAtranscription is paramount to functional mitochondrial biogenesis, we subsequentlymeasured the transcripts encoding for proteins that control mtDNA transcription,integrity and replication; namely mitochondrial transcription factor A (TFAM) andDNA polymerase γ (POLγ). Transcripts for TFAM (Tfam) were reduced in DOX-treated rats; however, the expression of POLγ (Polg) was unaffected (Fig. 3.2A).Increases in mitochondrial mass also involve the proliferation of mitochondrialmembranes, which in turn requires de novo synthesis of cardiolipin, the mainfunctional phospholipid of the mitochondrial inner membrane. Tafazzin (TAZ) isa transacylase required for cardiolipin metabolism that has been implicated in anumber of mitochondrial disorders that culminate in cardiomyopathy (Schlameand Ren, 2006). DOX treatment induced a small, although statistically significantdecrease in TAZ (Taz) mRNA levels (Fig. 3.2A).

A possible consequence of TFAM downregulation is a decrease in mtDNAquality and quantity. Indeed, in addition to its role as a mitochondrial transcriptionfactor, TFAM is also a major component of the nucleoid – a mitochondrialmaintenance structure (Campbell et al., 2012). Moreover, steady-state levels ofTFAM and mtDNA display a strong correlation (Campbell et al., 2012). Wetherefore assessed the copy number and integrity of mtDNA – two factors that fallunder the category of mitoepigenetics. ∗ Subchronic DOX treatment significantlydecreased the copy numbers of the mitochondrial COX1 gene per nuclear β2Mgene in heart but not liver tissue (Fig. 3.2C), suggesting a decreased mtDNAcopy number in the heart. There was no significant difference in the integrity ofmtDNA extracted from heart, suggesting that there is an equal extent, if any, ofmtDNA single- and double-strand break damage between both groups (Fig. 3.2B).

Finally, as a possible outcome of the above mentioned effects, we evaluatedtranscripts for the various components of the OxPhos machinery encoded by boththe nuclear and the mitochondrial genomes. We observed a general trend towardsa decrease in most of the transcripts assessed, most notably in two nDNA-encoded

∗Mitoepigenetics is an umbrella term for a group of factors that affect the expression of themitochondrial transcriptome, both nuclear and mitochondrial encoded (Ferreira et al., 2015).

3.2 Mitochondrial biogenesis 31

PGC-

1α

NRF-1

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Figure 3.2. DOX compromises mitochondrial biogenesis transcriptomics and mi-toepigenomics. (A, E) Heart mRNA levels of (A) proteins involved in the regulationof mitochondrial maintenance and biogenesis and (E) nuclear and mitochondrial genesencoding OxPhos subunits were quantified by qPCR and normalized to 18 S, TBP andβ-actin (n = 4–5 per group). (B) mtDNA integrity was examined by a PCR-basedassay as described (n = 5 per group). (C) mtDNA copy number was measured byqPCR (n = 3–5 per group). (D) Representative gels with the 10 kb and 121 bp prod-ucts. ∗p < 0.05 and ∗∗p < 0.01 for the indicated comparisons. Data are presented asmeans ± SEM.

32 Results

complex III subunits (UQCRC1 [Uqcrc1 ] and UQCRFS1 [Uqcrfs1 ]), one mtDNA-encoded complex I subunit (ND1 ) and one mtDNA-encoded complex IV subunit(COX1 ). Curiously, there was a trend towards upregulation of Cyt b (Mt-cyb)mRNA, similarly observed by us in an almost identical subchronic toxicity model(Pereira et al., submitted).

3.3 DOX alters acetyl CoA metabolism and trans-port transcripts and changes mitochondrialprotein acetylation patterns

Subchronic DOX treatment has previously been reported to decrease the expres-sion of proteins involved in the fatty acid β-oxidation pathway (Berthiaume andWallace, 2007b). To confirm this effect in our model, we measured the transcriptsfor PPARα, which regulates the transcriptional activation or repression of genesencoding for enzymes in FFA metabolism (Finck, 2007; Munday and Hemingway,1999), as well as transcripts for enzymes involved in fatty acid β-oxidation. Weobserved a marked decrease in PPARα (Ppara) and for both of the assessed β-oxidation enzymes (ACOX1 [Acox1 ], ACAA2 [Acaa2 ]) transcripts. Interestingly,transcripts for acetyl-CoA carboxylase 2 (ACC2 [Acacb]), which is repressed byPPARα (Munday and Hemingway, 1999), was also found to be reduced. Tran-scripts for CACT (Scl25a20 ), but not ACL (Acly) was also decreased, suggestingthat carnitine-mediated transport of acetyl moieties between the mitochondrialand cytosolic compartments may be impaired.

To test whether these findings were relevant for protein acetylation levelsand patterns, we carried out Western blot analyses of Ac-Lys in three differentsubcellular fractions – nuclear, mitochondrial and cytosolic –, so as to increase thespatial resolution of our findings. Total protein acetylation levels were unchangedin all of the three assessed subcellular fractions (Fig. 3.3B–D). Protein acetylationpatterns, however, were found to be altered in mitochondrial fractions. SAL-treated rats displayed inconsistent levels of a ≈ 63 kDa acetylated protein band,which was found to be present in three out of five animals. In DOX-treated rats,however, this band was completely absent (Fig. 3.3C).

3.4 DOX alters the transcription and activity ofhistone modulators

Histone modification by HATs and HDACs is one of the major mechanisms oftranscriptional regulation. The levels of these enzymes not only dictate the steady-state equilibrium of histone acetylation, but also determine the availability ofthese factors to be recruited to transcriptional regulatory complexes (Kouzarides,2007). Because of their role in modulating the epigenome, we next verified whethertranscripts from genes encoding HATs (CBP [Crebbp], Ep300, PCAF [Pcaf ]) andHDACs (HDAC1–10 [Hdac1–10 ]) were altered by DOX treatment. Two of thethree HATs mRNAs assessed, CBP and Ep300, were significantly downregulated

3.4 Histone modulators 33

245180135

100

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A

Figure 3.3. DOX alters the transcripts of genes involved in the metabolism andavailability of Ac-CoA in cardiac tissue, but does not change total lysine acetylationpatterns. (A) Heart mRNA levels of genes involved in the metabolism and transportof acetyl units were quantified by qPCR and normalized to 18 S, TBP and β-actin(n = 4–5). (B–D) Representative immunoblots (top) and summarized data (bottom) ofwestern blot experiments showing relative levels of proteins acetylated at Lys residues of(B) nuclear fractions, (C) mitochondrial fractions and (D) cytosolic fractions extractedfrom cardiac tissue. Numbers on the left side of blots denote regions that were deemedas quantifiable by densitometry analysis. ∗p < 0.05 and ∗∗p < 0.01 for the indicatedcomparisons. Data are presented as means ± SEM.

34 Results

Heart Liver0

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Figure 3.4. DOX alters transcripts and activity of histone modifiers. (A) Heart mRNAlevels of genes involved in histone acetylation and deacetylation were quantified by qPCRand normalized to 18 S, TBP and β-actin (n = 4–5). (B, C) The activities of (B) HATsand (C) HDACs were measured using commercial kits as described (n = 4–5). ∗p < 0.05and ∗∗p < 0.01 for the indicated comparisons. Data are presented as means ± SEM.

in DOX-treated animals (Fig. 3.4A). The majority of the targeted transcriptsencoding for HDACs displayed a trend towards downregulation, with a notableexception of HDAC1 (Hdac1 ), which was found to be increased in DOX-treatedanimals (Fig. 3.4A).

We next investigated the activities of these two sets of enzymes. Histoneacetyltransferase activity was found to be unaffected by DOX treatment in bothheart and liver tissue; however, HDAC activity was increased in heart tissue(Fig. 3.4B, C).

3.5 DOX alters SAM metabolism transcripts anddecreases global DNA m5C levels in cardiactissue

Maintenance of DNA methylation patterns is likely to depend on the levels ofDNMT1 and its required substrate SAM, the synthesis of which requires the

3.5 Global DNA m5C levels 35

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Figure 3.5. DOX decreases transcripts involved in the metabolism of SAM anddecreases global m5C content in cardiac tissue. (A) Heart mRNA levels of genesinvolved in DNA methylation and SAM metabolism were quantified by qPCR andnormalized to 18 S, TBP and β-actin (n = 4–5 per group). (B) Global m5C contentin DNA extracted from rat heart and liver tissue was examined by an ELISA assay(n = 5–6 per group). ∗p < 0.05 for the indicated comparisons. Data are presented asmeans ± SEM.

enzyme MAT2a (Lu and Mato, 2012). Subchronic DOX treatment reducedMAT2a (Mat2a) transcripts, thus suggesting an impairment in the synthesis ofSAM. However, levels of DNMT1 (Dnmt1 ) were unaltered after DOX treatment(Fig. 3.5A).

To assess whether DOX treatment affects DNA methylation in both heartand liver tissue, total DNA extracted from both tissues was analyzed with a m5CELISA-based assay. DOX treatment resulted in a significant decrease in DNAmethylation in heart but not in liver tissue (Fig. 3.5B).

Chapter 4

Discussion

Mitochondrionopathy is a well established early feature of DOX treatment (Wal-lace, 2003), however its role in the delayed, late-onset nature of DOX-inducedcardiotoxicity remains elusive (Berthiaume and Wallace, 2007b). It is not yetclear whether the decline in cardiac metabolic function is due to compromisedmitochondrial integrity and function, or rather due to a rewiring of the cardiacmetabolic profile via events upstream from mitochondria. The results from ourstudy lend credibility to our hypothesis that chronologically links both of theaforementioned events. Specifically, we propose that DOX initially disturbs the mi-tochondrial-dependent production of the main acetyl and methyl donors (Ac-CoAand SAM, respectively) and subsequently imprints a long lasting toxic epigeneticmemory that manifests itself through an aberrant metabolic transcriptome. Hereinwe provide a description of the initial events relevant to out hypothesis that aretaking place shortly after subchronic DOX treatment.

4.1 Mitochondrial biogenesisAs previously mentioned, disruption of mitochondrial function and biogenesis iswidely acknowledged as a key component in DOX-induced cardiotoxicity (Octaviaet al., 2012). Indeed, strategies aimed at increasing mitochondrial biogenesis, suchas treatment with resveratrol, endurance training and stimulation of the CO/hemeoxygenase (CO/HO) system, have proven to be effective in diminishing cardiacDOX toxicity (Ascensao et al., 2005; Suliman et al., 2007; Danz et al., 2009).Mitochondrial function is heavily dependent on a coordinated transcriptionalregulation of both the nuclear and the mitochondrial genomes (Scarpulla, 2006).The coactivator PGC-1α has emerged as a key component in the orchestration ofmitochondrial biogenesis and is highly expressed in tissues with high oxidativeactivity such as the heart (Ventura-Clapier et al., 2008). PGC-1α is a coactivatorof the transcription factors NRF-1 and -2 that in turn promote the expression ofTFAM, thus establishing a concerted activation of the nuclear and mitochondrialgenomes. In the present study, we demonstrate that subchronic DOX treatmentsignificantly decreases PGC-1α transcripts, as well as of its downstream targetTFAM. Our attempts at performing western blots for these proteins were notsuccessful; however, previous reports in the literature show that DOX decreases the

37

38 Discussion

content of both PGC-1α and TFAM, albeit using different treatment protocols thanour own (Suliman et al., 2007; Guo et al., 2014; Che et al., 2015). Nevertheless,decreased levels of PGC-1α and TFAM would account for the depression innuclear- and mitochondrial-encoded OxPhos transcripts in DOX-treated rats thatwe observed.

Another key index of mitochondrial biogenesis, or lack thereof, is cellu-lar mtDNA content. Our results show that DOX treatment decreases themtDNA:nDNA copy number in cardiac tissue, which may also be implicatedin the observed depression of mitochondrial-encoded OxPhos transcripts. In-terestingly, we did not observe reduced mtDNA:nDNA copy numbers in livertissue, suggesting that mtDNA alterations play a role in the cardiac-specific natureof DOX toxicity. Mitochondrial DNA strand-break damage, however, was notdetected at our assessed time point. This result is not surprising, since the kineticsof the mtDNA repair machinery are likely to be able to fully cope with the ratesof damage production that may be occurring at the time point we assessed. Forexample, in studies measuring the effect of ischemia–reperfusion in heart, knownto induce an instantaneous burst of ROS (Ambrosio et al., 1991), mtDNA damageis evident after 10 min of reperfusion, but returns to baseline levels after 60 min ofreperfusion (Bliksoen et al., 2015). Assuming, however, that DOX induced mtDNAstrand-break damage throughout the duration of our experimental protocol, onemay speculate that a possible mechanism to resolve single- and double-strandbreaks would be the complete elimination of the damaged molecule, thus account-ing, at least in part, for the observed depletion in mtDNA copy number. It isalso important to note, however, that the assay we employed is not sensible tospecific types of DNA damage that do not hamper DNA polymerase activity, suchas oxidized bases. Therefore, although we have determined that subchronicallyDOX-treated rats do not display mtDNA strand-break lesions, mtDNA damage inthe form of oxidized bases (such as 8-hydroxydeoxyguanosine [8-OHdG]) is likelyto be in place, as it has already been implicated in subchronic DOX treatment(Serrano et al., 1999).

Curiously, the above results partly account for an interesting observation in avery relevant study by Lebrecht et al. (2003). The authors, employing a delayedmodel of subchronic DOX toxicity in Wistar rats (sacrificed 31 weeks after the lastinjection), demonstrated a persistent 50 % decrease in mtDNA:nDNA copy numberand a selective downregulation of mitochondrial OxPhos transcripts vs. nuclearOxPhos transcripts. Consistently, they further demonstrated a selective decreasein the enzymatic activity of cytochrome c oxidase (COX, a multisubunit complexencoded both by nDNA and mtDNA) vs. the activity of succinate dehydroge-nase (SDH, encoded entirely by nDNA), again hinting at a mitochondrial-specifictranscriptional imbalance. This suggests that at later stages, the heart is ableto reestablish proper nDNA-encoded mitochondrial transcript levels, but notmtDNA-encoded ones. Future studies addressing this issue should be aimed atconfirming not only this observation but also additional nDNA-encoded mitochon-drial transcript levels, in order to provide a definitive answer to the unilateralnuclear reestablishment of mitochondrial transcription.

4.2 Metabolism 39

4.2 MetabolismAcetyl CoA

An additional major contributing factor to DOX-induced cardiotoxicity is themetabolic remodeling of cardiac tissue. We confirmed in our model previousresults that demonstrate a decrease in transcripts involved in fatty acid oxidationin hearts from rats treated subchronically with DOX (Berthiaume and Wallace,2007b). Importantly, this was not accompanied by an upregulation of ACC2,which would be expected in a programmed, well concerted metabolic shift sinceit encodes for an enzyme in an opposing pathway. This may be a contributingfactor to the inhibition of fatty acid oxidation associated with DOX cardiotoxicity(Abdel-aleem et al., 1997; Bordoni et al., 1999).

The available literature suggests an interesting relationship between fatty acidoxidation and Ac-CoA production. There is substantial evidence implicatingthe carnitine shuttle system in the acquired metabolic inflexibility. For instance,Carvalho et al. (2010) showed that octanoate, a medium-chain fatty acid that doesnot require protein-mediated import into the mitochondrion, is equally oxidized bycontrol and DOX treated rats. Conversely, long-chain fatty acids (which do requireprotein-mediated import) were shown to be inefficiently oxidized, suggesting thatfatty acid transport is impaired with DOX treatment. However, no differences ingene expression or protein content of carnitine palmitoyltransferase I B (CPT1B)were found in this model (Berthiaume and Wallace, 2007b; Carvalho et al., 2010).In a study investigating the effect of l-carnitine (the cofactor of CPT1B) infatty acid oxidation, Abdel-aleem et al. (1997) showed that the impairment inlong-chain FFA oxidation is restored in DOX-treated rats co-administered l-carnitine. However, the same animals also displayed an improvement in octanoateoxidation, suggesting that l-carnitine administration may be exerting its effectson β-oxidation itself rather than on FFA transport. Indeed, taking into accountthat DOX greatly increases the Ac-CoA/CoA-SH ratio in isolated rat heartmitochondria (Ashour et al., 2012), it is plausible that l-carnitine acts as anAc-CoA buffer, thus fully or partially restoring this equilibrium and thereby liftingthe Ac-CoA-imposed inhibition of 3-ketoacyl-CoA thiolase – the final enzyme ofthe β-oxidation cycle. More importantly for our present work, however, is thepossibility of maintaining normal histone acetylation dynamics by preventing theincrease in Ac-CoA/CoA-SH, thus potentially ameliorating the aberrant metabolictranscriptional profile. Our results support the role of l-carnitine in bufferingexcess mitochondrial Ac-CoA, since the observed downregulation of CACT mayfurther contribute to this metabolic instability.

Determination of the Ac-CoA/CoA-SH ratio could not be concluded at thetime of this writing; however, we assessed the cellular acetylome in advance soas to posteriorly correlate this parameter with the steady-state levels of the Ac-CoA/CoA-SH pair. No differences were found in whole cell levels of acetylatedproteins. However, acetylation of a ≈ 63 kDa protein was found to be absentin DOX-treated rats. Such a selective pattern of acetylation clearly indicatesthat in our model, whole cell protein lysine acetylation status is not limited byeventual alterations to the metabolome but is instead responding to cellular signals

40 Discussion

that may culminate in, for example, changes in protein synthesis, degradation,translocation, repression of acetylases or activation of deacetylases.

Future work is necessary to identify the protein corresponding to the absentband. At this point in time, however, we suspect that the protein in question maybe a member of the PGC-1 family for a number of reasons. First, both PGC-1αand PGC-1β are proteins subjected to dynamic acetylation and deacetylationin response to metabolic cues – particularly mediated by mitochondrial sirtuins(SIRT1, 3 and 5) (Kong et al., 2010; Canto et al., 2009; Kelly et al., 2009; Huanget al., 2010). Second, PGC-1α has been reported to colocalize in all of the threecellular compartments we assessed (nucleus, mitochondria and cytosol) (Wrightet al., 2007; Safdar et al., 2011). Indeed, although PGC-1α is known to lack thecanonical N-terminal mitochondrial targeting signal, it has been suggested toinstead contain cryptic internal signals (Safdar et al., 2011). Third, PGC-1α isknown to exist in multiple isoforms of different molecular weights in humans andmice (Ruas et al., 2012), meaning that we cannot discard the possibility of thisband corresponding to PGC-1α based on its migration to a non-canonical PGC-1αmolecular weight region (91–110 kDa) alone. Furthermore, the accuracy of ourchosen protein marker may also be put to question. Finally, and most importantly,our failed attempts at immunoblotting PGC-1α in nuclear and mitochondrialfractions displayed the same pattern observed in close proximity to the ≈ 63 kDaregion of the Ac-Lys immunoblot – that is, one single band in nuclear fractionsand two bands in mitochondrial fractions (Fig. A.1). Indeed, PGC-1α is frequentlyreported as a double band protein in Western blot experiments (Safdar et al.,2011; Miller et al., 2012), with some authors even distinguishing between the lightand heavy bands as PGC-1α and β, respectively (Garcia-Gimenez et al., 2011;Pohjoismaeki et al., 2012). Because in the PGC-1 immunoblot we did not observethe absence of the higher molecular weight band in mitochondrial fractions of DOX-treated rats, if this protein turns out to belong in fact to the PGC-1 family, thenwe would be confirming for the first time a selective mitochondrial deacetylationof a PGC-1 protein upon subchronic DOX treatment. We are currently workingon confirming this suspicion by means of PGC-1 immunoprecipitation followed byanti Ac-Lys immunoblotting.

S-adenosyl-L-methionine

Because SAM levels are sensitive to environmental cues such as diet (Poirieret al., 2001), a strong mitochondrial toxicant such as DOX is likely to alter SAMlevels as well. We are currently working on determining SAM and SAH levels bymeans of HPLC, but again this task is still unfinished. Nevertheless, we showthat MAT2a (the enzyme that catalyzes the last step in SAM synthesis) wasdownregulated, suggesting that SAM synthesis may be impaired. A reductionin the SAM/SAH ratio would imply a reduction in DNA methylase activity, notonly due to the limited substrate availability but also because SAH itself is apotent inhibitor of DNMTs (Selhub and Miller, 1992). Indeed, metabolic clearanceof SAH is a key metabolic determinant to multiple methyltransferase reactions(Selhub and Miller, 1992). Furthermore, MAT2a has been shown to colocalize inthe nucleus, specifically in chromatin-associated complexes. Absence of MAT2a

4.3 Epigenetics 41

in these complexes may further compromise the local production of SAM formethyltransferases reactions (Katoh et al., 2011), which may be more meaningfulthan global SAM synthesis for the purposes of DNA methylation.

4.3 EpigeneticsWe report for the first time that DOX treatment disturbs mRNA levels of HATsand HDACs, two important classes of histone modulators. While there wereno detectable differences in HAT enzymatic activity on either heart or livertissue, the increased activity of HDACs seen in cardiac tissue appears to correlatewith the only transcript that displayed increased levels (HDAC1 ). This resultsuggests that the expression of HDAC proteins may be following a similar trendto the one observed in the transcription patterns. The class I HDAC familyconsists of HDAC1, 2, 3 and 8. These are the most significant candidates forepigenetic modulation, since they colocalize predominantly in the nucleus andpossess high enzymatic activity toward histone substrates (Haberland et al., 2009).Members of the class IIa HDAC family (HDAC4, 5, 7 and 9), conversely, possessa very low catalytic activity (Fischle et al., 2002; Jones et al., 2008) and theirrepressive role has been attributed mostly to their function as a scaffold protein inmultiprotein transcriptional repressor complexes rather than any actual catalysis(Zhang et al., 2001). ∗ Thus, although we observe a generalized decrease in HDACtranscripts, our enzymatic activity assay is in agreement with a potential increasedexpression of class I HDAC proteins that would in turn be in concordance with themeasured transcripts. To establish a correlation between this result and epigeneticmodulation we intend to perform Western blot experiments against acetylatedhistones.

Global levels of m5C in DNA from DOX-treated rats were found to be decreasedin heart, but not liver tissue, also suggesting a relation with the cardiac-specificnature of DOX toxicity. Paradoxically, reduction in global DNA m5C contentwas accompanied by a depression in the transcript levels of multiple genes. DNAmethylation is an epigenetic mark mostly associated with transcriptional repression,therefore it is to be expected that a decrease in DNA methylation is followedby an increase in gene transcription. However, it is important to mention thatDNA methylation is but one element in the myriad of complexity layers coveringchromatin modification. In addition, the method of methylation quantificationemployed in this study was limited to a global analysis, and provides no informationregarding gene- or region-specific differences.

Theoretically, reduced DNA methylation would be in agreement with a poten-tial deregulation of folate metabolism and a concomitant decrease in the SAM/SAHratio, as less methyl donors would be available to participate in DNA methylation.However, the available literature suggests that the relationship between SAMavailability and DNA methylation does not follow this linear chain of thought.One in vitro study investigating the effect of folic acid supplementation in DNA

∗We will leave the class IIb HDAC family (HDAC6 and 10) out of this discussion sinceHDAC6 is only described as a cytoplasmic cytoskeletal deacetylase (Zhang et al., 2008) and therole of HDAC10 is currently unknown (Guardiola and Yao, 2002; Haberland et al., 2009).

42 Discussion

methylation shows that deprivation of folic acid decreases the SAM/SAH ratio but,also paradoxically, increases global DNA methylation (Farias et al., 2015). Thiseffect appeared to be mediated mostly by changes in the expression of DNMTs 1and 3a (which decreased by 50 % and increased by 80 %, respectively) rather thanvariations in the metabolome. To fully elucidate the mechanisms by which DOXdecreases DNA methylation, it will be essential to measure not only SAM andSAH levels but also the protein content of the different DNA methylases.

Chapter 5

Conclusions

In summary, the present study provides new data and confirms previous datademonstrating the occurrence of metabolic and epigenetic insults as well astranscriptome disruption shortly after cessation of subchronic DOX treatment.These results are in agreement with the hypothesis that metabolite-mediatedmanipulation of the cardiac epigenome is an underlying factor in the persistentnature of DOX-induced cardiotoxicity. Our study provides essential substantiatingevidence needed to motivate more ambitious follow-up experiments to furtherunravel the validity of this hypothesis using more thorough in vitro models andmore expensive in vivo delayed toxicity models.

Further complementary results are needed to fully allow for the speculationof a cause and effect relationship between the initial mitochondrial injury andthe also initial, albeit potentially permanent, epigenetic disruption. It will benecessary to verify if DOX-treatment disrupts the Ac-CoA/CoA-SH ratio inour experimental model and, subsequently, if the histone acetylome changesaccordingly. Likewise, the SAM/SAH ratio as well as the DNA methylasesprotein levels will also have to be assessed in order to compare with the observeddecrease in global DNA methylation. Results allowing for the establishment ofthe aforementioned relationship would motivate new avenues of research usingprimary cardiomyocytes so as to conclusively answer our experimental question.It would be interesting to verify if strategies that rescue mitochondrial functionin DOX models would prevent the concurrent epigenetic and transcriptionalalterations. Perhaps more interestingly, strategies using acetyl- and methyl-donorsupplementation or pharmacological manipulation of epigenetic enzymes wouldalso shed light on our question.

43

Appendix A

Supplementary Information

75

63

48

35

PGC-1α

SAL

DOXNuclear fraction

100

75

63

48

PGC-1α

SAL

DOXMitochondrial fraction

Figure A.1. Representative immunoblots of Western blot experiments showing relativelevels of PGC-1α in nuclear fractions and mitochondrial fractions extracted from cardiactissue. Western blots were carried out as described in Section 2.11 using a PGC-1antibody (1:1000; Santa Cruz Biotechnology; sc-13067).

45

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